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Page 1: Encyclopedia of Physical Science and Technology Molecular Biology
Page 2: Encyclopedia of Physical Science and Technology Molecular Biology

Table of Contents (Subject Area: Molecular Biology)

Article Authors Pages in the Encyclopedia

Cell Death (Apoptosis) Masato Enari Pages 541-554

Chromatin Structure and Modification

Fyodor D. Urnov and Alan P. Wolffe Pages 809-829

DNA Testing in Forensic Science Moses S. Schanfield Pages 589-602

Gene Expression, Regulation of Göran Akusjärvi Pages 501-516

Immunology-Autoimmunity K. Michael Pollard and Eng M. Tan Pages 679-691

Ribozymes Alessandra Poggi and John J. Rossi Pages 253-261

Translation of RNA to Protein R. A. Cox and H. R. V. Arnstein Pages 31-51

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Cell Death(Apoptosis)Masato EnariImperial College School of Medicine at St. Mary’s

I. OverviewII. Death Factors and Their ReceptorsIII. Apoptotic Proteases, CaspasesIV. Signal Transduction of Death

Factor-Mediated ApoptosisV. Cell-Free System in Apoptosis

VI. Apoptotic DNase, CAD and Its Inhibitor, ICADVII. Molecular Mechanism of CAD and ICAD

VIII. Physiological DNA Fragmentation andPhagocytosis of Apoptotic Cells

IX. Perspectives


Apoptosis The typical process in physiological cell deaththat is accompanied by nuclear and cytoplasmic con-densation, fragmentation of cell bodies, chromosomalDNA fragmentation, loss of mitochondrial function,and alterations of cell membrane composition. It isdistinct in these regards from necrosis. The term wascreated by Wyllie and Kerr.

Caspases Cysteine proteases, some of which are acti-vated during apoptosis. Some caspases are involvedin processing of cytokines.

Cell-free system A biochemical technique to be recon-stituted cellular events as in vitro reaction.

Death factors Proteins belonging to the tumor necrosisfactor (TNF) superfamily that occur on the cell sur-faces as membrane-bound factors and in the extracel-lular compartment as soluble factors. They kill target

cells expressing their corresponding receptors or to at-tenuate killing activity.

DNases Enzymes possessing DNA-cleaving activity.Some DNases participate in chromosomal DNA degra-dation during apoptosis.

DNA fragmentation Chromosomal DNA from apoptoticcells gives rise to a ladder pattern on agarose gels, dueto multimeric nucleosomal units (∼180 base pairs).

Death receptors Proteins belonging to TNF receptorfamily that occur on cell surface, and mediate killingby effector cells expressing their cognate ligands.

Intracellular signal transduction The cellular machin-ery that mediates external signals including hor-mones, neurotransmitters, cell growth, differentiationand death factors, or stress to their ultimate targets.

Phagocytosis The process by which phagocytes, such asmacrophages and neutrophils, engulf useless


and un-necessary cells.

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542 Cell Death (Apoptosis)

APOPTOSIS, or programed cell death, plays an impor-tant role in the development of organisms and in the main-tenance of homeostasis. The failure of apoptotic programscauses various diseases. So far, many genes regulatingapoptosis have been identified and the molecular mecha-nism of apoptosis is being clarified. In this article, I willdiscuss cell death elicited through death receptors.


Homeostasis in multicellular organisms is based on a bal-ance between life and death of cells. Apoptosis was recog-nized as a phenomenon distinct from necrosis by Wyllie

FIGURE 1 (a) Fas-induced apoptosis in lymphoid cells (WR19L cells overexpressing Fas). The cells were incubatedwith 0.5 µg/ml of an agonistic anti-Fas antibody at 37◦C for 120 min and their ultrastructure was examined undera transmission electron microscope. The electron micrograph of untreated cells is shown in the upper panel. Bars,1 µm. (b) Chromosomal DNA of growing cells (lane 1) or dying cells (lane 2) was run through a 1.5% agarose gel. Mindicates molecular weight markers.

and Kerr in 1972. In the necrotic process, swelling of cellsprecedes their explosion and results in the release of in-tracellular components that may be toxic to other cells.In apoptosis, the dying cells exhibit nuclear and cytoplas-mic condensation, fragmentation of cell bodies, chromo-somal DNA fragmentation into nucleosomal units, lossof mitochondrial function, and alterations of cell mem-brane composition (Fig. 1). Subsequently, apoptotic cellsare engulfed by phagocytes and neighboring cells, and arerecycled. Most cells suffering physiological cell death un-dergo the apoptotic process and the superfluous or harmfulcells generated during the developmental process are re-moved by apoptosis. For example, apoptosis occurs in tailresorption, neuronal network formation, clonal deletion of

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Cell Death (Apoptosis) 543

immature and autoreactive T cells, Wolffian and Mullerianduct regression during sexual development, tumor regres-sion, and in elimination of virus-infected cells. Further-more, it has been suggested that apoptosis occurs in manydiseases such as cancer, fulminant hepatitis, acquired im-mune deficiency syndrome (AIDS), diabetes mellitus, andneurodegenerative disorders such as Alzheimer’s diseaseand prion disease.

Growth and differentiation of cells are strictly regulatedby factors such as cytokines and low-molecular-weightcompounds such as steroid hormones. These factors aregenerally bound to the corresponding receptors in order totransduce the appropriate cell signals, to promote growthand differentiation. On the other hand, apoptotic cell deathis aggressively controlled by a number of polypeptides,so-called death factors exposed at the cell surface or cir-culating in the body as soluble factors in some situations.Growth and differentiation factors act via transcriptionalregulation through the activation of a series of protein ki-nases. On the other hand, death factors execute apoptosisthrough the activation of caspases, and many proteins es-sential for cell survival are degraded by these activated cas-pases. It is currently believed that apoptotic death is due tothe degradation of many functional proteins by caspases.Cell-free systems for the study of apoptosis have beenestablished and facilitate our understanding of the under-lying molecular mechanisms. Several factors involved inapoptotic pathways have been identified and part of thesepathways has been revealed by biochemical approachesbased on cell-free apoptosis system. Regulatory mecha-nisms for apoptosis include: the CED-3/caspase familyproteases and CED-4/Apaf-1 family which act as execu-tors of apoptosis; CED-9/Bcl-2 family (including anti-apoptotic and pro-apoptotic factors) which act as regu-lators of apoptosis, and many factors which contributeto apoptotic morphological changes have been identified.Here, I shall focus on the currently proposed molecularmechanisms of death receptor activity and caspase activa-tion. Moreover, I will also discuss the DNase responsiblefor apoptotic DNA fragmentation, both in vitro and invivo, and finally the mechanism by which apoptotic cellsare cleared.


Many cells have death receptors on the surface of theirplasma membranes and apoptosis is triggered by their cog-nate ligands. Death receptors belong to the superfamilyof tumor necrosis factor (TNF) receptors. Most consistof a cysteine-rich extracellular domain, a membrane-spanning domain, and a cytoplasmic domain containing

a death domain (DD) that is required for transducingapoptotic signals to cells. Six DD-containing death re-ceptors, namely Fas/Apo1/CD95, type I TNF recep-tor (TNFRI), DR3/Apo3/WSL-1/TRAMP, DR4/TRAIL-R1 (TNF-related apoptosis-inducing ligand receptor-1),DR5/TRAIL-R2/TRICK2/KILLER and DR6 have beenidentified so far. In addition, there are DD-less death re-ceptors such as type II TNF receptor (TNFRII), CD27,CD30, CD40, and lymphotoxin-β receptor. Among these,TNFRII, CD27, and CD30 induce the expression of bothTNF and TNFRI by their cognate ligands, and subsequentinduction of apoptosis appears to occur.

On the other hand, death factors are a member ofthe TNF family and exist on the cell surface or assoluble factors. Most death factors are synthesized astype II-membrane proteins and subjected to shedding bymembrane-associated metalloproteinases in order to gen-erate soluble forms of death factors. A well-establishedmechanism for shedding of death factors is seen in the caseof TNF. Membrane-bound TNF (memTNF) is cleaved atthe outer cellular membrane by a membrane-spanning pro-tease, so-called TNF α-converting enzyme (TACE), whichis a member of metalloproteinase-disintegrin (ADAM)family. memTNF is superior to soluble TNF (sTNF) inactivating TNFRII in various cellular responses, includ-ing T-cell proliferation, inflammation, and cytotoxicity.These results imply that memTNF regulates cellular re-sponses via restricted cell-to-cell interaction under physi-ological conditions and that sTNF may attenuate TNFRII-mediated responses. The shedding mechanism for otherdeath factors may be similar to that for TNF. Like in shed-ding of TNF, the membrane form of Fas ligand (mem-FasL) is also cleaved by an unknown metalloproteinaseother than TACE present on the plasma membranes. Inaddition, the soluble form of Fas ligand (sFasL) inhibitsmFasL-mediated apoptosis in human peripheral blood Tlymphocytes in vitro.

It is believed that death receptors are activated byligand-induced trimerization, as opposed to the activa-tion of growth factor receptors, which occurs by dimer-ization. Most death factors, but not all, are present astrimers. X-ray structural analyses have revealed that TNF-α, TNF-β, CD40L, and TRAIL are homotrimeric proteins.Among these, TRAIL has unique characteristics. TRAILrequires a zinc ion for biological activity and selectivelyinduces apoptosis in mouse tumor cells but not in nor-mal cells. Moreover, administration of soluble TRAIL tomice implanted with human tumors causes effective re-duction of tumor size without any injury of normal tis-sues. These results suggest that TRAIL may be applicableas an anti-cancer drug. However, a recent paper reportedthat TRAIL induces apoptosis in human hepatocytes,indicating that substantial liver toxicity might result if

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TRAIL were used in human cancer therapy. Before it canbe used in a clinical setting, it would be necessary to under-stand the molecular mechanisms by which TRAIL trans-duces signals from its receptor in human hepatocytes.


In the execution phase of apoptosis, caspase proteases areactivated by various apoptotic stimuli. Caspases belongto the cysteine protease family and were originally iden-tified as homologues of the ced-3 (cell death abnormal)gene product (an executor of apoptosis in Caenorhabdi-tis elegans). Indeed, most caspases induce apoptosis ifthey are overexpressed in growing cells and cell death canbe blocked by caspase-specific inhibitors. Therefore, cas-pases are accepted as executors for apoptosis in mammalsand this pathway appears to be conserved between species.

Caspases are synthesized as zymogens and the activeenzymes are generated by proteolytic cleavage of thesecaspase precursors. X-ray crystallographic studies haveshown that active caspases are heterotetramers consist-ing of two large subunits (∼20 kDa) and two smaller(∼10 kDa) subunits. So far, 14 caspases have been clonedfrom mammalian sources and divided into three sub-groups based on primary structure, phylogenetic analy-sis, and substrate specificity. Caspases cleave proteins onthe carboxyl side of aspartic acid with sequence speci-ficity. For example, caspase-1 shows preferential cleav-age at Asp in the Tyr–Val–Ala–Asp sequence, whereascaspase-3 cleaves at latter Asp in the Asp–Glu–Val–Asp.Caspases are classified as initiator, effector and cytokine-releasing caspases, according to their functions. Initiatorcaspases include caspase-2, -8, -9, and -10. These havelarge prodomains, which interact with specific adaptermolecules to convert the precursor to the active form. Ef-fector caspases, including caspase-3, -6, -7, and -14, haveshort prodomains and are activated by active initiator cas-pases. This regulatory mechanism is known as a proteasecascade. Other caspases found in mammals appear to serveas cytokine-releasing enzymes.

Gene disruption experiments have revealed the phys-iological functions of individual caspases in vivo. (1)Caspase-3- and -9-deficient mice show embryonic lethalphenotypes with neuronal hyperplasia, indicating thatcaspase-3 and -9 play an important role in neu-ronal development. Moreover, their embryonic fibrob-lasts are resistant to staurosporine-, etoposide-, UV-,and dexamethasone-induced apoptosis but not to Fas-mediated apoptosis. (2) Caspase-8-null mice are embry-onic lethal and established cells from these mice are re-sistant to cell death through death receptors includingFas, DR3, and TNFRI, whereas these cells are sensitive

to staurosporine-induced apoptosis. (3) Caspase-2 is re-quired for the formation of female germ cells and loss ofthe caspase-2 gene renders B cells resistant to granzymeB-induced cell death. (4) Patients suffering from au-toimmune lymphoproliferative syndrome (ALPS) typeII, which display immune regulatory defects, have mis-sense mutations in caspase-10. Caspase-10 mutants haveless caspase activity and block Fas- and TRAILR1-mediated apoptotic pathways. (5) Both caspase-1−/− andcaspase-11−/− mice are resistant to endotoxic shock,such as results from LPS treatment, and their geneproducts are required for the production of cytokines:caspase-1 is needed for secretion of IL-1α and caspase-11for secretion of IL-1α and β. As discussed above, eachcaspase is important for both various developmental andpathological processes. Knockout analyses for other cas-pase genes are still under investigation and will revealindividual in-vivo functions for each caspase in the nearfuture.


As described earlier, Fas- or TNFRI-mediated apoptosis istriggered by binding of the cognate death ligands (Fig. 2).In general, apoptosis induced by most death factors canstill occur in the presence of inhibitors of protein or RNAsynthesis, suggesting that the death factor-mediated apop-totic processes proceed without de novo synthesis of eitherproteins or RNAs unlike the processes involved in growthand differentiation, and that all of components for apop-totic signaling are constitutively present in cells. There isan essential domain required for transduction of the deathsignals into cells, so-called death domain (DD), foundin the cytoplasmic region of both Fas and TNFRI. Im-munoprecipitation analyses have suggested that a proteincomplex, designated as the death-inducing signaling com-plex (DISC), is recruited to the cytoplasmic domain of Fasfollowing the interaction of Fas with Fas ligand (Fig. 2).Using the yeast-two hybrid technique with the cytoplas-mic region of Fas or TNFRI as baits, several moleculesthat specifically bind to the cytoplasmic region of thesereceptors have been discovered. FADD (Fas-associatingprotein with death domain)/MORT-1 is a small adapterprotein with a molecular mass of 26 kDa and a death do-main at the its C-terminus. FADD is recruited to trimer-ized cytoplasmic region of Fas and binds to Fas via in-teractions between the death domains (Fig. 2). Nuclearmagnetic resonance (NMR) studies have shown that thedeath domain of Fas consists of six antiparallel, amphi-pathic α-helices arranged in a novel fold. Because thereare many charged groups from amino acids on the protein

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FIGURE 2 Signal transduction of Fas- and TNFRI-mediated apoptosis.

surface, the binding between death domains may be me-diated by ionic interactions. Deletion mutant experimentshave shown that the N-terminal region of FADD but notits death domain is necessary for transducing death sig-nals. In addition, the FADD with deletion of the N ter-minus works as a dominant-negative mutant against theFas-mediated system. Therefore, this region has beendesignated as death effector domain (DED). Similarly,the DD-possessing protein, TRADD (TNFRI-associateddeath domain protein), has been found as an adaptermolecule that specifically binds to TNFRI. Unlike FADD,TRADD does not have DED, but is essential for mediatingapoptosis induced by TNFRI. Subsequently, it has beenshown that TRADD binds to FADD via DD interactions(Fig. 2). That is, TRADD is recruited to the cytoplasmic re-gion of trimerized TNFRI that consequently binds FADD.Thus, Fas and TNFRI utilize the same transducer and sharethe apoptotic machinery downstream of FADD. The sig-nal through TNFRI is more complex than that of Fas be-cause of the diversity of signals from TNFRI. For example,RIP (receptor-interacting protein), originally identified asa Fas-binding protein with DD motif, preferentially binds

to TRADD and is involved in the activation of transcrip-tional factor NF-κB that is responsible for stimulating theproliferation of thymocytes (Fig. 2). Although RIP has akinase domain at the N terminus, this kinase domain is dis-pensable for activating NF-κB. Several analyses indicatethat NF-κB appears to protect against TNFRI-mediatedapoptosis via up-regulation of c-IAP2 (cellular inhibitorof apoptosis protein 2). Up-regulated c-IAP2 appears tobind TRAF2 (TNF receptor-associated factor 2), whichcan bind RIP and inhibit apoptotic signaling from TNFRI(Fig. 2). Both negative and positive apoptotic signals fromTNFRI might be necessary for fine tuning the decision todie or not.

Two independent approaches involving the yeast two-hybrid experiments with DED of FADD as a bait and pu-rification of factors binding to the cytoplasmic region ofFas, have revealed that procaspase-8 binds to the DED.Procaspase-8 contains two DED motifs at the N terminusthrough which it binds to FADD and a caspase homol-ogous region at the C terminus (Fig. 2). Indeed, DISCcontains both FADD and procaspase-8, and recruitedprocaspase-8 is processed to the active enzyme near the

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inner plasma membrane. Several lines of evidences sug-gest that procaspase-8 may be proteolytically processedto the active form by simple oligomerization. However,the precise mechanism is still unclear and may requireunknown additional factor(s) that may be in DISC or onthe inner plasma membrane, may be required to gener-ate active caspase-8 more efficiently. Once caspase-8 isactivated in cells, it cleaves effector procaspases locateddownstream of it such as caspase-3, -6, and -7 in order toamplify the apoptotic signal (Fig. 2). Finally, cells showa variety of apoptotic features such as described above,due to cleavage of more than 100 substrate proteins whichcontributes to DNA fragmentation, loss of mitochondrialfunction, and the maintenance of nuclear structure andplasma membrane.

An alternative apoptotic pathway via mitochondriahas been found by the in vitro reconstitution assay (de-scribed in section V). Based on a dATP/ATP-induciblecell-free system, three factors responsible for process-ing procaspase-3 have been purified and identified. Theseinclude Apaf-1, which is found as a mammalian homo-logue of CED-4 known to be another executor of apop-tosis in C . elegans, cytochrome c (Apaf-2) that is mainlypresent in the mitochondrial intermembrane space, andprocaspase-9 (Apaf-3). Cytochrome c is released fromthe intermembrane space of mitochondria into the cy-toplasm during apoptosis induced by a variety of apop-totic stimuli, including DNA-damaging agents, protein ki-nase inhibitors (staurosporine), and death receptors (Fig.3). Moreover, the release of cytochrome c is blockedby anti-apoptotic proteins belonging to the Bcl-2 fam-ily such as Bcl-2 and Bcl-xL (Fig. 3). Recent obser-vations have indicated that cytochrome c-deficient miceshow an embryonic lethal phenotype with defects in ox-idative phosphorylation. In addition, their embryonic fi-broblasts are resistant to stresses such as UV irradia-tion, serum withdrawal and staurosporine. The mech-anism by which cytochrome c is released from mito-chondria by apoptotic stimuli remains elusive, althoughsome hypotheses, including opening of a specific chan-nel for cytochrome c, alteration of the permeabilitytransition pore (PTP) that regulates inner mitochondrialmembrane potential, or swelling and subsequent rup-ture of the outer mitochondrial membrane, have beenproposed.

Apaf-1 possesses a region homologous to theprocaspase-prodomain, known as the caspase-recruitingdomain (CARD) at the N terminus, a region homolo-gous to CED-4 in the middle part and WD-40 repeatstructure that appears to be involved in protein–proteininteractions. The released cytochrome c interacts withtwo cytosolic proteins, Apaf-1 and procaspase-9, anddATP/ATP in the cytoplasm to form a complex known as

the apoptosome (Fig. 3). Apaf-1 binds to procaspase-9via CARD motifs and procaspase-9 is processed usingboth Apaf-1 and energy from dATP/ATP hydrolysis toconvert procaspase-9 to the active form. Cytochrome cappears to play a role in overcoming the inhibition of theApaf-1 active site masked by the WD-40 repeat. Onceactivated, effector caspases downstream of caspase-9 aresequentially activated (Fig. 3).

It has recently been shown that there are two cell typeswith different sensitivity to Fas signaling induced by anagonistic anti-Fas antibody. In type I cells, procaspase-8 israpidly activated following receptor engagement, whereasin type II cells the activation of procaspase-8 is delayed,although both type I and type II cells show similar kineticsof Fas-mediated apoptosis and loss of mitochondrial func-tion. In addition, Bcl-2 inhibits apoptosis in type II but nottype I cells. Why do these cells die in a similar mannerto each other regardless of the amount of active caspase-8and yet show different blocking activity by Bcl-2? Oneexplanation is that there are two distinct pathways in Fas-mediated apoptosis, a direct pathway from caspase-8 toeffector caspase in type I cells and a caspase-8-mediatedmitochondrial pathway in type II cells. It is likely thatBcl-2 is only able to block the latter pathway. However,this hypothesis is still controversial in view of the reportthat physiological Fas ligand but not an agonistic antibodykills both types of cells similarly regardless of the Bcl-2expression level. These results might depend on the effi-ciency of precise trimerization of Fas. The evidence thatthere are two distinct pathways in the Fas system has comefrom the analyses of Bid-deficient mice. If Bid, a Bcl-2family member, is cleaved by caspase-8, truncated Bid(tBid) can translocate from the cytosol to mitochondria,subsequently, cytochrome c is released, executing apop-tosis. Administration of an agonistic anti-Fas antibody towild-type mice in vivo causes death with hepatocellularapoptosis and haemorrhagic necrosis in the liver within 3hours, whereas Bid-deficient mice survive treatment withthis antibody. In addition, hepatocytes from Bid−/− miceare resistant to this antibody in vitro, suggesting that thetBid-mediated pathway via mitochondria predominantlyworks in hepatocellular apoptosis induced by agonisticFas antibody. It will be interesting to determine whetheror not Apaf-1−/− and caspase-9−/−conditional knockoutmice (because of the embryonic lethal phenotype) showa similar response as Bid-deficient mice to the adminis-tration of the anti-Fas antibody. To investigate the actualsignaling pathways activated by the physiological Fas lig-and in vivo, it will be necessary to administer soluble Fasligand to Bid-deficient mice.

Some death receptors also activate caspase proteasesby ligation with their cognate ligands. Like in TNFRI sig-naling, DD regions of activated death receptors including

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FIGURE 3 Apoptotic pathway via mitochondria.

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DR3 and TRAILR2 recruit DISC (that consists of FADDand procaspase-8) via the DD of TRADD. Unlike Fas,TNFRI, and DR3, TRAILR1 can transmit apoptotic sig-nals and activate caspase-8 in FADD-deficient fibroblastcells. This suggests that TRAILR1-mediated apoptosisis caspase-8-dependent and FADD-independent, althoughan unidentified FADD-like adapter molecules cannot beruled out at present. The response mediated by DR6 hasnot, as yet, been well characterized.

In conclusion, ligand-induced trimerized death re-ceptors activate an apical protease, procaspase-8, viaFADD or an unidentified functional homologue. Acti-vated caspase-8 can sequentially activate effector cas-pases responsible for cleaving a variety of death substratessuch as ICAD/DFF45 (discussed below), lamin, fodrin,and poly(ADP-ribose) polymerase. In addition to the di-rect pathway from initiator caspases to effector caspasesthrough Fas, active caspase-8 can transduce death signalsinto mitochondria through the translocation of caspase-8-cleaved Bid from cytosol to mitochondria. Uptake of tBidinto mitochondria promotes the release of cytochrome cto the cytosol by unknown mechanisms. Once cytochromec is released, it can generate active caspase-9 via forma-tion of the apoptosome. Thus, death receptors appear touse two different pathways for particular tissues in somesituations. The two different pathways ensure the deathsignals are amplified and target cells killed.


The establishment of cell-free systems has provided uswith detailed insight into the cognate molecular mech-anisms of various cellular functions, including gen-eral/specific transcriptional regulations, RNA editing,protein synthesis and protein degradation, and overallmetabolic pathways. The well-established biochemicalhallmark of apoptosis is chromosomal DNA degrada-tion resulting in multimers of 180 bp-nucleosomal units.Neither protein- nor RNA-synthesis inhibitors block Fas-mediated apoptosis, suggesting that all of the componentsfor apoptotic induction through Fas are present in cells inlatent forms.

A cell-free system for apoptosis was first establishedby exposing isolated nuclei to various extracts andmonitoring nucleosomal DNA fragmentation. That is,when isolated nuclei from healthy cells were treatedwith extracts from dying cells (but not from growingcells), they showed apoptotic features, including nuclearmorphology with peripheral condensation of chromatinand nucleosomal DNA fragmentation. This cell-freesystem can be reproduced using the extracts from cellssubjected to different apoptotic stimuli, including UV

radiation and treatment with ceramide (N-acyl-erythro-sphingosine). In addition, extracts from cells activated byFas in the presence of a caspase inhibitor do not causeDNA fragmentation in the cell-free reaction. Indeed,extracts from proliferating cells, in the presence ofrecombinant active caspase-3, induce nuclear apoptosis,indicating that the factors responsible for apoptotic DNAfragmentation are downstream of the caspase cascade.It emerged that these factors form a complex composedof heterodimeric proteins of caspase-activated DNase(CAD)/DNA fragmentation factor 40 (DFF40)/caspase-activated nuclease (CPAN) and an inhibitor of CAD(ICAD)/DFF45 (discussed in the following).

As described above, if dATP is mixed with extracts fromproliferating cells, the extracts have the ability to inducenuclear apoptosis mediated by caspase-3 activation in acell-free reaction. Using this system, Wang’s group havepurified the factors responsible for processing procaspase-3 and identified three gene products, namely Apaf-1,cytochrome c and procaspase-9. Thus, biochemical ap-proaches using cell-free systems for apoptosis has demon-strated several important aspects in the field of apoptosis.

During apoptosis, biochemical procedures have fre-quently shown loss of mitochondrial function. In order toclarify the regulations of mitochondrial apoptosis, isolatedmitochondria have been used. During the early stages ofstudy in this field, the function of mitochondria in apop-totic processes was overlooked due to no apparent changeson morphology. However, the observations that Bcl-2 (ananti-apoptotic protein) is abundant in the mitochondria,and that apoptotic processes are often accompanied bya decrease in the mitochondrial membrane potential andby mitochondrial swelling led to a more detailed inves-tigation of mitochondrial function during apoptosis. Fur-thermore, apoptogenic factors, such as cytochrome c andapoptosis-inducing factor (AIF) that had been discoveredas a nuclear apoptosis-inducing flavoprotein using a cell-free system, are released from the intermembrane spaceof mitochondria into the cytosol to induce apoptosis. Re-cently, another factor secreted from mitochondria duringapoptosis, called Diablo/Smac, has been reported. Thisfactor suppresses the functions of proteins belonging tothe inhibitor-of-apoptosis (IAP) family, inhibiting the pro-tease activity of caspases including caspase-3, -7 and -9.Cell death is likely to be accelerated as a result of inhibitionof IAP. Thus, there appear to be various steps to ensure cellkilling. How is the loss of mitochondrial function inducedby the various apoptotic stimuli? For example, Bax (Bcl-2-associated X protein), which belongs to a member ofthe pro-apoptotic Bcl-2 family, induces apoptosis. Whenadded individually to purified mitochondria-free cytosol,neither mitochondria nor Bax can individually induce theactivation of caspases that lead to nuclear apoptosis. In

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contrast, the addition of both Bax and mitochondria trig-gers the release of cytochrome c from mitochondria to cy-tosol, which activates the caspase cascade. In addition, ithas been shown that Bax directly triggers cytochrome c re-lease from mitochondria in a cell-free reaction. It has beenproposed that Bax may release cytochrome c from mito-chondria by modulating voltage-dependent anion chan-nel activity and/or causing limited permeabilization of themitochondrial outer membrane although this mechanismis still controversial. The apoptotic signals from variousstimuli mediated by mitochondria should be investigated,dissected, and reproduced by biochemical approaches us-ing a cell-free system described here.


A biochemical hallmark of apoptosis is chromosomalDNA degradation. It was early on proposed that one ofseveral known enzymes, including DNase I, DNase II,DNase γ , and cyclophilins, contribute to chromosomalDNA fragmentation during apoptosis, although the molec-ular mechanism understanding apoptotic DNA fragmen-tation by these DNases was unclear.

Most apoptotic stimuli commit the cells to apoptosisthrough the activation of the caspase cascade. As describedin section V, several groups, including our own, have es-tablished a caspase-3-inducible cell-free system, in whichnot only chromosomal DNA in nuclei but also naked plas-mid DNA is cleaved by caspase-3-activated cell extracts,and used it to identify a responsible factor(s), designated asCAD, for apoptotic DNA fragmentation. We purified CADfrom lymphoid cells using the caspase-3-inducible cell-free system. We also noticed that an inhibiting-factor(s),designated as ICAD, is present in the extracts from non-apoptotic cells and identified ICAD as a 32-kDa protein.Molecular cloning of ICAD has revealed that long-form(331 amino acids; ICAD-L) and short-form (265 aminoacids; ICAD-S) of ICAD, which are generated throughalternative splicing of the same messenger RNA (Fig. 4a).Independently, other groups have purified a latent formof CAD consisting of heterodimers (DFF40 and DFF45)from human HeLa cells or an active form of CAD namedCPAN from human Jurkat cells, using a system similar toours.

Both ICAD-L and -S (ICAD) are acidic proteins withisoelectric points (pI) of around 4.5. A homology searchhas shown that ICAD-L has high similarity to humanDFF45, suggesting that mouse ICAD-L is a counterpartof human DFF45. ICAD-L is ubiquitously expressedin a variety of tissues at the same level as ICAD-S.ICAD carry two caspase-3-cleavage sites corresponding

to amino acids at 117 (DEPD117) and 224 (DAVD224)(Fig. 4a). Analyses of point mutations at two sites have in-dicated that the cleavage of ICAD at Asp117 and Asp224by caspase-3 is crucial for the inactivation of ICAD func-tion. Interestingly, the inhibitory activity of the ICADwith single point mutations at both sites was completelyresistant to treatment with caspase-3. Among the cas-pases 1 to 8, caspase-7 also cleaves ICAD in an in vitroassay. However, active caspase-7 translocates from cy-tosol to mitochondrial and microsomal fractions, whereasactive caspase-3 is still present in cytosol during Fas-induced apoptosis in mouse liver in vivo, suggestingthat ICAD in cytoplasm and nuclei are mainly cleavedby caspase-3. This is consistent with the observationthat caspase-3-deficient cells undergo apoptosis withoutchromosomal DNA fragmentation, although caspase-7 isactivated.

ICAD are thermostable at 90◦C for 5 min and resistantto denaturants such as 6 M guanidium hydrochloride, 8 Murea and 0.1% SDS. ICAD specifically inhibit the CADactivity, but not DNase I and DNase II activities, by bind-ing to it. In addition, ICAD can completely inhibit nuclearapoptosis induced by extracts from Fas-activated cells inthe presence of an inhibitor of caspase-3. Furthermore,overexpression of caspase-3-resistant ICAD mutants sup-presses apoptotic DNA degradation induced by diverseapoptotic stimuli including death factors, growth factorstarvation, anti-cancer drugs, and γ -irradiation, indicatingthat CAD needs to be activated to degrade chromosomalDNA in many apoptotic situations.

Molecular cloning of CAD has revealed that the mouseCAD gene encodes a basic protein of 344-amino acids anda pI of 9.7. Mouse CAD comprises 14 cysteine residues;methionine and cysteine residues at positions 1 and 2, re-spectively, are removed during the maturation of the pro-tein (Fig. 4b). Human CAD is composed of 338 aminoacids and is highly homologous to mouse CAD, with anidentity of 75.9% between both primary structures. Elevencysteine residues, most of which exist as reduced thiolgroups, are conserved between mouse and human CAD.The C terminal region of CAD contains a stretch of basicamino acids with the features of a nuclear localization sig-nal (Fig. 4b). Active CAD is a very unstable protein andeasily aggregates under nonreducing condition, in contrastto the CAD/ICAD-L complex. The stability of CAD is en-hanced by addition of reducing reagents such as dithiothre-itol (DTT) and reduced glutathione, suggesting that inter-and/or intra-molecular crosslinking of free thiols in CADmay cause the aggregation and inactivation of CAD func-tion. Some DNases are inactivated by reagents that modifyfree thiols, such as iodoacetamide and N-ethyl maleimide,whereas CAD activity is not inhibited by such reagents,suggesting that free thiols in CAD are not required for its

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FIGURE 4 Structure of mouse CAD and ICAD.

activity. In addition, it is known that DNA increases thestability of DNase I. Therefore, it would be interesting todetermine whether free-thiol-modifying reagents and/oraddition of DNA could enhance CAD stability and facili-tate the structural analyses.

Various endonucleases are implicated in apoptotic DNAdegradation, including nonmetal ion-requiring, Mg2+-requiring, Mg2+- and Ca2+-requiring, and Mg2+- orCa2+- requiring DNases. CAD belongs to the class ofMg2+-requiring endonucleases. The optimal pH for CADactivity is around 7.5. CAD activity is inhibited by Zn2+,in accordance with the observation that Zn2+ inhibitsapoptotic DNA degradation in some intact cells. CADdoes not digest single-stranded DNA or RNA. It cleavesdouble-stranded DNA leaving 5′-phosphate and 3′-hydroxyl ends; consistent with the fact that the DNAfragments generated during apoptosis are susceptible toend labeling by deoxynucleotidyl transferase (TUNELreaction). Furthermore, there is no consensus CAD-cleavage sequence on either genomic or plasmid DNA,but CAD appears to preferentially cleave sequences withrotational symmetry (5′-R, R, R, Y ⇓ R, Y, Y, Y-3′) orwith AT-rich region. In addition, CAD cleaves chromatinbetter than micrococcal nuclease. So far, micrococcal nu-clease has been used for chromatin research to investigate

chromatin functions but CAD might be a useful tool forsuch analyses.

Homology search has shown that CAD has no appar-ent overall similarity with any proteins. It has, however,been found that CAD shares a N-terminal region con-sisting of about 80 amino acids, so-called CAD or celldeath-inducing DFF45-like effector (CIDE) domain, withother proteins, including mouse and human ICAD, mouseFsp27, human and mouse CIDE-A, mouse CIDE-B, andDrosophila melanogaster DREP-1, based on databasesearches. Structural analyses have shown that the CADdomains from CAD and CIDE-B have a fold of α/β

roll type with a novel protein-protein-binding motif, andthat ICADs and CAD appear to interact through CADdomains.


In the absence of ICAD-L, CAD is expressed in an in-soluble form in various host cells including Escherichiacoli (E. coli), insect cells and mammalian cells. How-ever, coexpression of CAD and ICAD-L dramatically en-hances CAD activity induced by active caspase-3, and

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recombinant CAD is recovered in the cytosolic fractionas a CAD/ICAD-L complex. This process is reproducedusing an in vitro coupled transcription and translation sys-tem. Newly synthesized CAD aggregates in the absenceof ICAD-L in the in vitro reaction, and functional CADis produced only in the presence of ICAD-L, although theexpression level of CAD is the same whether or not ICAD-L is present. These results suggest that ICAD-L works asa chaperone to help the correct folding of CAD. These re-sults have been confirmed by the finding that chaperone-like activity of ICAD-L is seen even in refolding of purifiedand denatured CAD from inclusion bodies from E. coli inthe presence of high concentration of reducing reagents.This process does not require ATP, whereas reticulocytelysates in combination with ICAD-L enhance a refoldingprocess for denatured CAD in an ATP-dependent man-ner. These results imply that one or more ATP-dependentenhancer may participate in the folding process of CADunder physiological conditions. General chaperone sys-tems including Hsp70 may function in this process. Invitro refolding studies should reveal which factor(s) is in-volved in the folding of CAD in the near future. Anal-yses of the functional differences between ICAD-L andICAD-S have revealed that ICAD-S has less chaperone-like activity than ICAD-L and mainly exists as a homo-oligomeric complex (oligomerization of ICAD is depen-dent on their concentration). The finding that ICAD-L ispredominantly complexed with CAD and that only ICAD-S is purified from our assay may be due to the differenceof chaperone-like activity between ICAD-L and ICAD-S.Thus, when CAD is newly synthesized, ICAD-L binds tothe nascent chain of CAD on the ribosome to suppressaggregation of CAD and to help proper folding (Fig. 5).ICAD-L is incorporated into a CAD/ICAD-L complex toinhibit CAD activity. Caspase-resistant ICAD-L is likely

FIGURE 5 Model for the function of CAD and ICAD in apoptosis.

to have chaperone-like activity since the aggregation ofCAD is suppressed by coexpression with it. Thus ICAD-Lworks as a double safeguard against dangerous CAD func-tion. Once caspase-3 is activated by an apoptotic stimulus,ICAD-L is cleaved and released from CAD. The releaseof ICAD-L from complex permits active CAD to concen-trate in nuclei and to degrade chromosomal DNA (Fig. 5).Mouse CAD lacking the nuclear localization signal at theC terminus, (consisting of amino acids position 3 to 329,with the 15 basic amino acids of CAD primary sequencedeleted), still has DNase activity, but it cannot induce DNAfragmentation in nuclei. These observations suggest thatthe C terminal basic region actually works as nuclear lo-calization signal and is not required for DNase activity.

It is thought that there are two steps in apoptotic DNAdegradation. At the initial stage of apoptosis, chromoso-mal DNA is cleaved into 50- to 300-kilobase pair (kb)-sizefragments, followed by the cleavage of large fragments tonucleosomal units. Cyclophilins have been proposed ascandidates for large chromosomal degradation. However,the expression of caspase-resistant ICAD in cells blocksnot only small-size nucleosomal degradation but alsolarge-size chromosomal degradation induced by apoptoticstimuli such as Fas and staurosporine, indicating that CADis responsible for both steps. Large-size fragments couldbe due to preferential cleavage at nuclear scaffolds withAT tracts by CAD. This is consistent with the fact that nolarge-size DNA degradation was detected in the thymo-cytes from ICAD-deficient mice during dexamethasone-,etoposide-, and staurosporine-induced apoptosis.

Apoptosis is accompanied by nuclear condensationas well. When active CAD is incubated with isolatednuclei, CAD itself has an ability to induce apoptoticmorphological changes with chromatin condensed aroundthe nuclear periphery. ICAD-deficient thymocytes also

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show no chromatin condensation in nuclei after apop-totic stimulation. On the other hand, significantly, dyingcells overexpressing caspase-resistant ICAD show apop-totic chromatin condensation without DNA fragmenta-tion. These results suggest that apoptotic chromatin con-densation is caused by CAD in particular tissues such asthymocytes and that another factor(s) is involved in thisprocess in some situations. We also cannot rule out thepossibility that CAD has dual functions, DNA fragmen-tation and condensation activities, because of completedenatured structure of CAD in ICAD-null cells (CAD ispresent as an insoluble form) but not transformants ex-pressing caspase-resistant ICAD (endogenous CAD andICAD are still present as a soluble form), or that cleavedICAD cooperatively work with cellular factor(s) to con-dense chromatin although cleaved ICAD themselves haveno chromatin condensation activity. Recently, it has beenreported that a protein other than CAD, called Acinus,is responsible for apoptotic chromatin condensation how-ever this remains to be confirmed.


Extensive DNA fragmentation and chromatin condensa-tion are hallmarks of apoptosis and are tightly regulatedby the CAD/ICAD system. It is thought that these pro-cesses play an important role in cell killing. To revealphysiological functions of the CAD/ICAD system in vivo,Zhang et al. have established ICAD-deficient mice. Thesemice develop normally compared to wild-type mice anddo not show any pathological signs. However, thymocytesfrom ICAD-null mice are obviously lacking CAD activityand show neither DNA fragmentation nor chromatin con-densation following exposure to apoptotic stimuli. Con-sistent with the observations described above, no func-tional CAD is expressed, at least in thymocytes, in theabsence of ICAD. Moreover, transgenic mice expressingcaspase-resistant ICAD-S (ICAD-Sdm; it is ubiquitouslyexpressed under the control of elongation factor 1 α pro-moter) have been established. ICAD-Sdm is expressedin the major organs of the transgenic mice, and isolatedthymocytes from the transgenic mice undergo apoptosiswithout DNA fragmentation after irradiation with γ -rays.However, these transgenic mice have no phenotypic ab-normality during development, like ICAD-deficient mice.The thymocytes from ICAD-null mice, but not from wild-type mice, are partially resistant to several apoptoticstimuli including dexamethasone, etoposide, and stau-rosporine, whereas thymocytes from ICAD-Sdm trans-

genic mice die as efficient by as those from wild-type micefollowing exposure to γ -rays. Why is there such a discrep-ancy even though there is no CAD function in thymocytesin both cases? There is a possibility that either endoge-neous ICAD cleaved by caspase-3 or CAD/ICAD-Sdmcomplex may contribute to some killing activity in thy-mocytes from ICAD-Sdm transgenic mice. ICAD mightbe required for amplification of death signal in some con-ditions, or the modification of anti-apoptotic proteins incells. It has recently been reported that ICAD-null mice ex-hibit enhanced spatial learning and memory accompaniedby the increase (∼8%) of granule cells in the hippocampaldentate gyrus region compared to wild-type mice. Detailanalyses would be elucidated whether or not CAD/ICADsystem is actually involved in the induction of cell death.

It has been thought that phagocytes recognize the apop-totic cells undergoing DNA fragmentation and ingest andrecycle the dead cells, and that cleaved DNA is utilizedas a marker for the recognition of apoptotic cells byphagocytes. However, phagocytosis of dying thymocytesfrom ICAD-Sdm transgenic mice, in which DNA degra-dation is not seen, occurs as efficiently in the case ofwild-type mice, indicating that DNA fragmentation is dis-pensable for phagocytosis. Moreover, it has recently beenobserved that TUNEL-positive cells are still present invarious tissues from ICAD-Sdm transgenic mice at simi-lar levels as in wild-type mice after γ -irradiation. Whendying cells overexpressing ICAD-Sdm are cocultured withmacrophages, their intact DNAs are quickly processed intonucleosomal units. Furthermore, reagents that block theacidification of lysosomes inhibit DNA degradation in-duced by macrophages, suggesting that lysosomal DNasein engulfing cells is involved in the generation of TUNEL-positive cells in various tissues in vivo. A candidate forthe responsible DNase during phagocytic DNA fragmen-tation is DNase II existing in lysosomes as an acidicDNase. The recent molecular cloning of nuc-1, whichis responsible for DNA degradation during programmedcell death in C . elegans, has revealed that the gene prod-uct is a homologue of mammalian DNase II. This reportproposes a model for apoptotic DNA degradation com-posed of three distinct steps (Fig. 6). The first step ap-pears to involve an unidentified endonuclease to produceTUNEL-positive DNA. The enzyme working in this stepwould be a CAD-like DNase. In the second step NUC-1converts TUNEL-positive to TUNEL-negative DNA. Inthe final step, “cell-corpse DNA” is completely digestedby engulfment-dependent nuclease(s). DNase II createsDNA fragments having 3′-phosphate ends rather than the3′-hydroxyl ends that are required as primers for termi-nal deoxynucleotidyl transferase. In the mammalian sys-tem, after cleavage of DNA by lysosomal DNase II, the3′-phosphate groups are presumably removed by acid

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phosphatase, which is abundant in lysosomes, giving riseto TUNEL-reactive ends. Figure 6 summarizes the modelproposed for apoptotic DNA degradation in the mam-malian system, based on the C. elegans paradigm: Oncecaspase-3 is activated by one of various apoptotic stimuli,ICAD is inactivated and releases from CAD. ActivatedCAD cleaves chromosomal DNA generating TUNEL-reactive ends. This is followed by DNase II-mediatedDNA degradation that gives rise to TUNEL-negative DNA(Fig. 6). Due to participates of lysosomal enzymes con-taining a large amount of acid phosphatase in mammalianphagocytes, TUNEL-reactive ends could be generatedin engulfing cells in mammals. Finally, complete diges-tion of cell-corpse DNA occurs during phagocytosis torelease and recycle free nucleotides (Fig. 6). Recently,a CAD/ICAD system has been identified in Drosophilamelanogaster. Thus, the mechanism for apoptotic DNAdegradation is likely to be conserved between mammals,flies, and worms. The detailed mechanism will be eluci-dated by further use of genetic and molecular biologicalapproaches.

A variety of factors contributing to engulfment havebeen reported in nematodes, flies and mammals. Someof the responsible factors, including CED-1, CED-2,CED-5, CED-6, CED-7, CED-10, and CED-12 in C.elegans, Croquemort (CRQ) in Drosophila melanogaster,and a phosphatidylserine receptor in mammals have re-cently been identified. The CED-6 protein contains aphosphotyrosine-binding (PTB) domain at its N terminusand putative Src-homology domain 3 (SH3)-binding sites.It has been shown that this protein restores the functionof the clearance of apoptotic cells in engulfment-defectiveced-6 mutants from C. elegans. The CED-7 protein is ho-mologous to ABC (ATP-binding cassette) transporters, is

FIGURE 6 Model for apoptotic chromosomal DNA degradation in vivo.

localized in the plasma membrane and the endoplasmicreticulum, and appears to function both in dying cellsand in engulfing cells during the engulfment process. Ithas been suggested that CED-7/ABC1 is required for thetranslocation of molecules that interact between the cellsurfaces of the apoptotic cells and phagocytes in both C. el-egans and mammals. The ced-5 gene product, which func-tions in cell-corpse engulfment in C. elegans, is similar tothe human DOCK180 and the Drosophila melanogasterMyoblast City (MBC). It has been suggested that bothDOCK180 and MBC are involved in the extension of cellsurfaces and may function during phagocytosis in differ-ent species. CED-2 and CED-10 are counterparts of thehuman adapter protein CrkII and the human GTPase Rac,respectively. These factors are likely to function in phago-cytes, but not in dying cells, to regulate phagocytosis.Based on genetic crossing experiments, CED-10 appearsto be downstream of CED-2 and CED-5 and GTPase sig-naling must regulate the polarized extension of cell sur-face for phagocytosis. Drosophila CRQ is a CD36-likereceptor expressed on the cell surface of macrophages;genetic analyses have shown that it is required for phago-cytosis of apoptotic corpses. Apoptosis is also accompa-nied by the exposure of phosphatidylserine on the outercell surface of plasma membranes. Phosphatidylserine hasalso been regarded as a target molecule for phagocytosis;the phosphatidylserine receptor has been cloned by phagedisplay technique. It is expressed on various cells includ-ing macrophages, fibroblasts, and epithelial cells, and isfound in databases of Drosophila melanogaster and C. el-egans as a protein of unknown function. An antagonisticantibody against the phosphatidylserine receptor blocksengulfment of dying B and T cells by phagocytes, indi-cating that this receptor is essential for phagocytosis of

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at least some apoptotic cells. Thus, although various fac-tors participating in phagocytosis have been identified, itwill be necessary to find further components involved inthis process to understand the molecular mechanism ofclearance of apoptotic cells in our bodies.


Not only cell growth and differentiation factors but alsodeath factors regulate homeostasis of organisms. Thedeath factors activate a caspase cascade through their spe-cific receptors. Although it is said that procaspase-8 is self-cleaved simply following activation by oligomerization,the precise mechanism is still unclear. Some unknownadditional factor(s) may be required for this process togenerate active caspase-8 more efficiently and remain tobe identified by future analyses. Furthermore, it is knownthat there is caspase-independent apoptosis. It would beimportant to investigate what kinds of factors are involvedin such apoptotic pathways. The mechanism of apoptoticDNA fragmentation has been largely clarified by biochem-ical approaches, but the mitochondrial pathway for deathsignals remains elusive. How are apoptotic signals trans-duced in mitochondria? Which factors participate in mi-tochondrial apoptosis? Cell-free systems using isolatedmitochondria may lead to a satisfactory solution of theseproblems.

Cell death occurs without cell-autonomous DNA degra-dation in cells devoid of CAD activity and developmentalprocesses appear to be normal (without drastic changes)despite the lack of functional CAD. Why has a compli-cated CAD/ICAD system evolved in our bodies? One pos-sibility is that this system precludes the transformation ofphagocytes by dangerous genes such as oncogenes or vi-ral genes. Another possibility is that the cell-autonomousDNA fragmentation mechanism reduces autoimmune re-sponses against nucleosomal DNA, which is known asa strong autoantigen. Nucleosomal DNA released fromapoptotic cells during developmental processes could leadto autoimmunity. Alternatively, the CAD/ICAD systemmay promote the recycling of the DNA components. Anal-yses of CAD/ICAD system-deficient mice may cast lighton these questions. Furthermore, the number of genes rec-ognized as being involved in phagocytosis of apoptoticcells is gradually increasing due to genetic and molecu-lar biological analyses. Thus, the understanding of detailmolecular mechanisms of apoptosis deepens as science isprogressed, so the elucidation of overall mechanisms forapoptosis through death receptors may not be so far off.


I thank all members of Dr. Nagata’s laboratory in Osaka BioscienceInstitute and Osaka University Medical School for helping during myinitial works. I am grateful to Drs. S. Nagata and H. Sakahira in OsakaUniversity Medical School for helpful comments on the manuscript andto Dr. C. Weissmann at the Imperial College School of Medicine forsupports, discussion, and encouragement.






Budihardjo, I., Oliver, H., Lutter, M., Luo, X., and Wang, X. (1999).“Biochemical pathways of caspase activation during apoptosis,” Annu.Rev. Cell Dev. Biol. 15, 269–290.

Bratton, S. B., MacFarlane, M., Cain, K., and Cohen, G. M. (2000).“Protein complexes activate distinct caspase cascades in death receptorand stress-induced apoptosis,” Exp. Cell Res. 256, 27–33.

Ferri, K. F., and Kroemer, G. (2000). “Control of apoptotic DNA degra-dation,” Nat. Cell Biol. 2, E63-4.

Green, D. R., and Reed, J. C. (1998). “Mitochondria and apoptosis,”Science 281, 1309–1312.

Liu, Q. A., and Hengartner, M. O. (1999). “The molecular mechanismof programmed cell death in C,” Elegans. Ann. N. Y. Acad. Sci. 887,92–104.

Nagata, S. (1997). “Apoptosis by death factor,” Cell 88, 355–365.Nagata, S. (2000). “Steering anti-cancer drugs away from the TRAIL,”

Nat. Med. 6, 502–503.Nagata, S. (2000). “Apoptotic DNA fragmentation,” Exp. Cell Res. 256,

12–18.Raff, M. (1998). “Cell suicide for beginners,” Nature 396, 119–122.Stroh, C., and Schulze-Osthoff, K. (1998). “Death by a thousand cuts:

An ever-increasing list of caspase substrates,” Cell Death Diff. 5, 997–1000.

Thornberry, N. A., and Lazebnik, Y. (1998). “Caspases: Enemies within,”Science 281, 1312–1316.

Yeh, W. C., Hakem, R., Woo, M., and Mak, T. W. (1999). “Gene targetingin the analysis of mammalian apoptosis and TNF receptor superfamilysignaling,” Immunol. Rev. 169, 283–302.

Wyllie, A. H., Kerr, J. F. R., and Currie, A. R. (1980). “Cell death: Thesignificance of apoptosis,” Int. Rev. Cytol. 68, 251–306.

Wyllie, A. H. (1980). “Glucocorticoid-induced thymocyte apoptosis isassociated with endogenous endonuclease activation,” Nature 284,555–556.

Zhang, J., Liu, X., Scherer, D. C., van Kaer, L., Wang, X., and Xu, M.(1998). “Resistance to DNA fragmentation and chromatin condensa-tion in mice lacking the DNA fragmentation factor 45,” Proc. Natl.Acad. Sci. U.S.A. 95, 12480–12485.

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Chromatin Structureand Modification

Fyodor D. UrnovAlan P. WolffeSangamo Biosciences

I. The 6-Billion ChallengeII. Chromatin: A Brief History of Scholarship

III. Chromatin Structure: The Histonesand the Nucleosome

IV. The Disruption and Modification of ChromatinStructure as a Tool to Control the Genome

V. Chromatin and Transcription: A Synthesis


Chromatin A complex between genomic DNA and pro-teins (mostly histones) that compacts the genome intothe eukaryotic nucleus and enables its functionality.

DNA methylation The covalent modification of cytosineto yield 5-methylcytosine. In vertebrates occurs ex-clusively within the context of a CpG dinucleotide.Produced by DNA methyltransferase and interpretedby dedicated DNA-binding proteins that effect tran-scriptional repression in part via recruiting histonedeacetylase.

HAT Histone acetyltransferase. Enzyme that catalyzesthe acetylation of the ε-NH2 group in the side chainof lysine residues in the core histones’ amino terminaltail. Commonly used in transcriptional activation.

HDAC Histone deacetylase. Enzyme that catalyzes theremoval of the acetyl residue from ε-acetyllysine in the

core histones’ amino terminal tail. Commonly used intranscriptional repression.

Histone Small (ca. 110 amino acids), highly basic pro-tein that is rich in lysine and arginine. Assumes a dis-tinctive 3-helix “histone fold” tertiary structure withan extended amino terminal tail. Four “core” his-tones (H2A, H2B, H3, H4) form an octamer thatwinds DNA onto itself to form the nucleosome coreparticle.

Nuclear hormone receptor (NHR) A transcription fac-tor whose activity is regulated by a small molecule(the ligand), such as a steroid (estrogen, cortisol, etc.)or an amino acid (thyroid hormone). Most NHRs aretranscriptional activators in the presence of ligand;some NHRs also act as transcriptional repressors inits absence.

Locus (pl. loci) Geneticists’ term for “specific locationon the chromosome (in the genome).”


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Nucleosome A union of eight core histones, one linkerhistone, and ca. 160 base pairs of DNA. The elementarybuilding block of the chromatin fiber.

Remodeling An ATP-dependent process of disruptinghistone–DNA contacts in chromatin. Requires the ac-tion of dedicated macromolecular machines such as theSWI/SNF complex. An integral component of gene ac-tivation and repression.

Transcription The process of RNA synthesis on a DNAtemplate. Catalyzed by RNA polymerase.

COMPACTION of the eukaryotic genome into the nu-cleus must occur such that the instructions in the DNAare accessible to molecular machines that effect replica-tion, transcription, and repair. This challenging task is per-formed by uniting the DNA with histone proteins to yieldchromatin—a dynamic, complex structure that controlsthe genome by both compacting and revealing it. Each180 base pairs of DNA is complexed with eight moleculesof core histone and one linker histone to yield the nucle-osome. A wide variety of modifications and alterationsof the nucleosomal fiber occur in vivo, including ATP-dependent disruption of histone-DNA contacts, and thecovalent modification of the histone tails by acetylation.The enzymatic machines that generate such modificationsare targeted to specific loci in the genome by various reg-ulators to effect gene activation and repression.

The enormously complex program of gene expres-sion that unfolds during ontogeny in eukaryotes is un-equivocally contingent on an accurate interpretation ofthe genome’s contents by transcriptional regulators. Theiraction is abetted by, and intimately functionally linkedto, the native structure assumed by their DNA target invivo: the nucleoprotein fiber of chromatin. This com-plex union of DNA with histone and other proteinsensures that the genome is accommodated within thenucleus by winding each 180 base pairs of DNA into anucleosome, and by subsequent higher-order folding ofthe nucleosomal fiber. Chromatin is not a passive scaffold,and undergoes a wide variety of structural transitions inresponse to developmental, environmental, and internalgene regulatory stimuli. A host of dedicated enzymaticcomplexes effect these transitions both in a genome-wide(i.e., after DNA replication) and a localized fashion, whenthey are recruited by transcriptional regulators to medi-ate gene actvation or repression at specific loci. Uniquepatterns of covalent post-translational histone modifica-tion, and perturbation of histone-DNA contacts or higher-order chromatin fiber structure that result from such tar-geting exert a potent regulatory effect on the underlyingDNA.


The functionality of the human genome—i.e., its capac-ity to control an extraordinarily complex program of celldivision, differentiation, and morphogenesis during em-bryonic development and adult ontogeny—is enabled inthe face of several formidable challenges.

The first one is topological: the combined length of asingle cell’s worth of human DNA is ca. 2.3 m (thereare ∼6.6 × 109 base pairs of DNA in the diploid humangenome and each base pair is 0.32 nm long) and this ex-traordinary quantity of nucleic acid—in the form of 46separate molecules—is packaged into that cell’s nucleus,i.e., a sphere with a diameter of ∼5 µM. Thus, the lengthof the DNA fiber (which is 2 nm wide) exceeds that of itsnatural container by a factor of ∼500,000.

The second challenge is that of the signal-to-noise ra-tio: laden with molecular atavisms and genomic parasites,the genome’s ∼800 Mb of genetic information (countingeach base pair as two bits, i.e., four base pairs per byte)make Finnegan’s Wake seem lucid and succinct by com-parison, since only ca. 20 Mb (∼3%) represent DNA thatcodes for protein. An even smaller fraction is “regulatoryDNA”—i.e., stretches of the genome that contain molec-ular instructions on how and when its ∼40,000 genes areto be activated and silenced. The exact percentage of thegenome assigned to such regulatory function is unknown,but a conservative estimate would be ca. 1% (8 Mb). Thus,the genome is a remarkably messy manual, in which adeluge of genetic gibberish (or, in less emphatic terms,of DNA to which we have not yet been able to assign afunction) hides the occasional useful instruction or snip-pet of data. The molecular complexes that have evolvedto interpret the genome—sequence-specific DNA bindingproteins and their cofactors—are thus challenged by a ma-jor problem of parsing the content of the genome in searchof relevant information.

As if this were not enough, the physicochemical prop-erties of DNA—a very long polymer—make its aqueoussolutions extraordinarily viscous (as anyone who has everworked with human genomic DNA knows from experi-ence, solutions of even 10 mg/ml cannot be pipetted if theDNA is not sheared first into fragments of smaller size).All interactions relevant to the biology of the genome,however, occur—and quite robustly—in an aqueous envi-ronment that contains ∼100 mg/ml DNA (6–8 pg in ca.65 femtoliters)!

All these challenges are very successfully met: in-side the nucleus, the DNA of the genome finds itself apowerful ally as a large number of dedicated molecularmachines assemble it into a highly structured fiber bywinding it around specialized proteins called “histones”;

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Chromatin Structure and Modification 811

the resulting entity—chromatin—presents the genomein compact, interpretable, and soluble form, and—mostimportantly—enables a remarkably rapid response to agreat variety of internal and external stimuli.


Cytological, genetic, biochemical, and biophysical stud-ies of the past 100 years have offered many insights intothe structure of chromatin and chromosomes and into themechanistic aspects of its role in enabling gene expressionand chromosome behavior. Our current notions of chro-matin structure and function offer interesting testimony tothe validity of T. Kuhn’s well-known thesis on the role of“paradigms” in scientific inquiry. The first half of the 20thcentury witnessed the protein constituents of the nucleusbeing awarded the role of carriers of genetic information,with DNA relegated to a minor role of a scaffold. After thediscovery of the nucleosome in 1973–1974, some 20 yearsof scholarship in eukaryotic transcription proceeded underthe auspices of notions developed in studying gene regu-lation in bacteria, and thus the histones underwent quitethe proverbial reversal of fortune and were considered arepressive, obstructive scaffold instead; bona fide tran-scriptional control was only thought to occur on stretchesof naked, histone-free DNA.

Following the genetic and biochemical characterizationof complex machines that modify chromatin in the mid-1990s, the past few years have offered a dramatic andemphatic shift in our appreciations of its intranuclear func-tion: no longer viewed as monotonous or merely repres-sive, chromatin is now considered an essential componentof most gene regulatory pathways in vivo. We now seethe nucleus as populated with enzymatic complexes thatremodel chromatin in a targeted fashion to achieve a widevariety of regulatory responses from the DNA. The currentview of transcriptional and genomic control, therefore, isone of a complex and mutually beneficial symbiosis be-tween the protein regulators of the genome and the nucle-oprotein architecture of chromatin.

The etymology of the term “chromatin” traces its ori-gins to cytological studies of the late 19th century, when“thread-like structures”—chromosomes—were revealedin dividing cells by staining with dyes. These were pre-sumed to consist of “chromatin” (a term proposed by W.Waldeyer in 1888) whose chemical nature was completelyobscure. Fortunately for science, the contemporaneousdiscovery by F. Miescher of DNA in human lympho-cytes eventually prompted an analysis into whether thesubstance of chromosomes is in any way related to thechemical entity extracted by Miescher from cells known

as “nuclein.” In 1889, work by R. Altmann demonstratedthat it consists of nucleic acid and protein. A functionalconnection between either of these two chemical sub-stances and the phenomenon of heredity remained elusive,however.

In the 1900–1920s, studies by T. H. Morgan and a dis-tinguished pleiade of his junior colleagues (C. B. Bridges,H. K. Muller, and A. H. Sturtevant) in the fruit flyDrosophila melanogaster physically placed genes ontochromosomes, connecting genetics and cytology. Bio-chemical analysis performed by A. Kossel in the 1890s–1910s led to the discovery of histones and protamines—the small, highly positively charged protein components ofchromosomes. Their apparent biochemical diversity, cou-pled with a notion of DNA as a chemically monotonousentity, prompted the belief into their role as carriers of ge-netic information. This theory was placed in very strongdoubt in 1944 by work of O. T. Avery, C. M. MacLeod,and M. McCarty at the Rockefeller Institute, and firmlylaid to rest in 1952 by A. D. Hershey and M. Chase at theCold Spring Harbor Laboratory.

After a combined effort by chemists and X-ray crys-tallographers, including P. Levene, A. Todd, E. Chargaff,R. E. Franklin, and M. H. F. Wilkins, made their indeli-ble contribution to the 1953 discovery by J. D. Watsonand F. H. C. Crick that DNA is a double helix, a 20-year avalanche of experimentation revealed the foundationof genetic information maintenance and transfer in livingsystems, mostly prokaryotes. This period witnessed threevery important studies that implicated chromatin in effect-ing eukaryotic gene regulation.

Cytological studies of mammalian cells have revealedan interesting gender bias: the interphase (i.e., nondi-viding) nuclei of female, but not male, cells containeda small, microscopically dense entity termed the “Barrbody.” In 1958, work from S. Ohno demonstrated thatthis entity corresponds to one of the two X chromosomesin the female karyotype. Soon after, in 1961, geneticexperiments by M. Lyon showed that the female genomeis functionally hemizygous for sex-linked loci—i.e., thatof the two X chromosomes in mammalian females, oneis genetically silent (this phenomenon is known as “Xchromosome inactivation”). The combination of thesedata provided evidence for a correlation between thestructure of specific chromosomes and their state of ac-tivity. In recent years, work from many laboratories, mostnotably those of S. Tilghman and R. Jaenisch, offeredseveral remarkable insights into the mechanistic aspectsof X chromosome inactivation (see below). Importantly,observations made by Ohno and Lyon extended earlier(1928) studies by E. Heitz on plant chromosomes, whichlead to the discovery that the contents of the nucleuscan be divided into heterochromatin and euchromatin,

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i.e., microscopically distinct compartments that appearedto represent chromosomes that condensed to distinctdegrees: heterochromatic domains in chromosomes werethus correctly deduced to contain silenced genes.

The complementary correlation between chromosomedecondensation and gene activation was provided by stud-ies in insects, whose salivary glands contain giant chro-mosomes that yield easily to microscopic examination. Aterminally differentiated tissue destined for destruction inthe metamorphosis from a larva to the adult insect, the sali-vary gland effects a very interesting solution to a demandfor high protein production: it replicates its DNA withoutintervening mitosis, and all the resulting DNA fibers foreach chromosome coalesce—side by side—into a macro-scopic body termed “the polytene chromosome.” Theirsize and certain other structural features have made theminvaluable model systems in biology. Most importantly,cytologic analysis of changes in chromosome structureduring larval development have revealed that at definedtimepoints, specific stretches of these giant chromosomesdecondense and form “puffs”—localized swellings. It wascorrectly deduced quite early on that such decondensationis related to the activation of genes that reside in thosestretches of the chromosome.

In 1960, H. Clever and U. Karlsson combined theirefforts of trying to determine, why the insect molting hor-mone, the steroid ecdysone, causes such dramatic mor-phological changes during insect development. The in-jection of purified ecdysone into larvae of the midgeChironomus lead to a premature and dramatic puffingof specific stretches of polytene chromosomes: thus, hor-mone action was for the first time connected to regulationof gene activity and, importantly, to a localized alteration(decondensation) of chromosome structure. A molecularmechanism, or correlate, of this striking phenomenon wasnot obtained until studies in 1984 by K. Zaret and K.Yamamoto, and subsequent experiments by T. Archer andG. Hager (Section IV.A).

Finally, 1964 saw the publication of a discovery whoseimpact resonated only in 1996, but quite emphatically:the observation by V. Allfrey, and A. E. Mirsky that hi-stone proteins are subjected to postranslational covalentmodification via the acetylation and methylation of ly-sine residues in their NH2-terminal tails (Sections IV.Aand IV.C). Because the modifications reduce the positivecharge of the histones (and thus have the potential to al-ter the way histones interact with DNA), it was immedi-ately suspected they might have regulatory consequences.Conclusive evidence to that effect was obtained in 1998(Section IV.C).

Inherent technical limitations of cytological and bio-chemical methods described could not illuminate themolecular structure of protein–DNA contacts within chro-

matin. The application of biophysical techniques in theearly 1970s led to significant progress in this field,and structural understanding improved accordingly, as a7 A X-ray crystal structure of the elementary subunit ofchromatin—the “nucleosome core particle” (146 bp ofDNA wrapped around the an octamer of core histones)—was described in 1984, and a 3.1 A resolution structure—in1991. Finally, in 1997, T. J. Richmond and colleagues pub-lished a 2.8 A nucleosome core particle structure—thus,we now understand the elementary composition and struc-ture of chromatin in very considerable detail. The nextsection describes the histone proteins, genetic evidencefor their role in regulating transcription, work in the 1970sthat led to the nucleosome hypothesis, and concludes witha description of the structure of the nucleosome.


A. The Histones

As discovered by Kossel early this century, the primaryprotein residents of the nucleus are small highly basic pro-teins called “the histones” (there are approximately 36 mil-lion nucleosomes, each containing nine histone moleculesand 180 bp of DNA, in the nucleus of a human cell). Onlyfive different histones are sufficient to assemble chro-matin: four core histones (H2A, H2B, H3, and H4; twoof each are present in each nucleosome) and one linkerhistone (H5; one per nucleosome). A technological Atlasfor molecular biology, gel electrophoresis through suchmatrices as polyacrylamide and agarose has been an in-valuable tool in studying the genome, and Fig. 1 shows adenaturing (i.e., performed in the presence of detergent,such as sodium dodecyl sulphate, SDS) polyacrylamidegel on which the histones are resolved.

The histones’ primary amino acid sequence offer sev-eral glimpses into their function: these proteins are small(between 102 and 130 amino acids long) and very rich inlysine or arginine. For example, of the 103 amino acids inhuman histone H4, 11 are lysine and 14 are arginine. Thishas an immediate electrostatic implication for histone be-haviour in vivo: the pKa values for these amino acids’ sidechains (10.0 and 12.0, respectively) indicate that at phys-iological pH, the average histone H4 molecule carries ca.24.99 positive charges on itself.

An interesting and informative aspect of histone biol-ogy is the extraordinary degree of sequence conservationbetween these proteins across taxa: histone H4 in humansand in tomato (Lycopersicon esculentum) is identical inlength and sequence with the exception of three highlyconservative substitutions (i.e., Val61 Ile). Thus, there

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FIGURE 1 The core histones (lane 1), histones H3 and H4, or thecore histones together with the linker histones (lane 3) resolvedon a polyacrylamide gel.

is considerable irony in the brief (ca. 1900–1952) histor-ical prominence that histones played as the putative ve-hicles of genetic data: DNA was incorrectly throught tobe monotonous in sequence, but histones far exceed theDNA in their invariance, both from nucleosome to nucleo-some, and between species. Such remarkable resistance tomutational pressure is particularly striking when one con-siders that—as discussed in the next section—only 75%of each histone is actually required to assemble a nucle-osome core particle. As shown in Fig. 2, all four corehistones can be assigned an identical secondary structure:the COOH-terminal 75% wind into 3 α-helices separatedby two loops (this portion of the histone functions in as-sembling the histone octamer and in arranging DNA ontoitself), while the structure of the NH2-terminal 25% (“thetail”) is not known. The conservation of the primary aminoacid sequence of the tail is strong evidence for its func-tional prominence, which is discussed below.

FIGURE 2 The core histones. The sequence of the amino terminal tails is indicated. The COOH-terminal domain isdepicted schematically at the bottom.

The “linker histone”—histone H1—is so named be-cause of its physical location in the chromatin fiber (seefollowing). It is slightly larger than the core histones (ca.20,000 Da), very rich in lysine, and assumes a very inter-esting structure shown in Fig. 3, called “the winged helix”(helix 3 is thought to contact the DNA). Several transcrip-tional regulators in metazoa are isomorphous to histoneH1, and the implications of this near-congruence fortheir physical location within chromatin and mechanismswhereby they affect gene expression are discussed here.

B. Experimental Evidence for the Histones’Role in Controlling Gene Expression

Before a discussion of chromatin structure details, it ishelpful to summarize in vivo evidence indicating that his-tones play a role in gene regulation. Simple a priori con-siderations suffice to realize that histones are required forcell viability—since chromosome condensation and sub-sequent segregation during mitosis is contingent on his-tones, cells would fail to divide in their absence and die,and this complicates genetic analysis (as eloquently putby one molecular biologist, “dead cells don’t tell stories”).An elegant solution was implemented by M. Grunstein, inwhose lab a strain of budding yeast, Saccharomyces cere-visiae, was engineered for such a study: the promoter ofthe histone H4 gene was replaced with one that could beinactivated with a simple change in growth medium (inthis case, the addition of glucose). The gradual depletionof histone H4 from chromatin does not lead to instant celllethality, and thus effects of “genetically dechromatinizingDNA” can be studied.

Use of such approaches by Grunstein’s lab, as well asof M. Osley, F. Winston, and M. Smith, offered a veryunexpected result: general notions of chromatin as re-pressive packing material would indicate that most ofthe genome should become spuriously active in the ab-sence of chromatin. Experimental observations indicated

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FIGURE 3 The linker histone—a ribbon representation of its“winged-helix” structure.

otherwise, however, and showed that histone depletion—and a concomitant decrease in the extent to which DNA isassembled into chromatin—does not have nucleus-widetranscriptional consequences, although the upregulationof specific genes was indeed observed.

Conclusive evidence to this effect was recently providedin work from the lab of R. Young. This study made useof the fact that the entire genome of budding yeast hasbeen sequenced, and it was therefore possible to performgenome-wide expression profiling analysis. This remark-able experiment requires a custom-made “microarray”: asilicon chip containing a grid, into each cell of which a dif-ferent single-stranded nucleic acid probe corresponding toa given yeast gene is placed and immobilized (a total of5900 genes were thus analyzed in one experiment). Mes-senger RNA is prepared from wild-type and mutant cellsat defined timepoints following inactivation of the histoneH4 gene, and the change in the levels of each message isdetermined by hybridizing each mRNA sample to a sep-arate chip (in actual fact, for detection purposes, a copyof the mRNA labeled with a fluorescent dye is prepared)and then measuring the difference—if any—in the levelsof the mRNA hybridizing to each cell in the array.

The major result from this study is that expressionof ca. 75% of the yeast genes is not significantly al-tered by histone depletion. This indicates that in buddingyeast, chromatin does not have a genome-wide transcrip-tional repression function. Remarkably, of the remain-ing 25% of the genes, ca. 15% were upregulated morethan threefold, while 10% were downregulated more thanthreefold. These data suggest that nucleosomes have gene-specific roles in transcriptional control, and, surprisingly,that the assembly into chromatin is not only required forthe repression of some genes but also for the activation ofsome others.

As elaborated in the next section, a examination of thenucleosome structure fails to explain these data—all nu-cleosomes appear to be the same by available structuralcriteria, and assemble the entire genome into what seemsto be a homogeneous fiber (see following). Remarkably,thinning out that fiber has markedly idiosyncratic effectson genome behavior—and some understanding as to pos-sible mechanisms for enabling such nonuniform responsesis beginning to emerge.

C. Ontology of the Nucleosome

Evidence that DNA in the nucleus may be organized intosome sort of repetitive entity first came from analysis ofthe way the genome inside the nucleus is seen by nucleases(i.e., enzymes that degrade DNA by cleaving the phospho-diester bond between two adjacent nucleotides), both en-dogenous to the cell and ectopic. Such experiments by R.Williamson in 1970, and by D. Hewish and L. Burgoyne in1973, demonstrated that DNA released from the genomein this way assumes a nonrandom length distribution—which was unexpected, because nucleases were not knownto have a substrate preference within DNA. By way of ex-ample, Fig. 4 shows an agarose gel containing DNA sam-ples after treatment with a nuclease. When the enzyme

FIGURE 4 Chromatin treatment with micrococcal nuclease(MNase) yields a nonrandom distribution of DNA fragments—evidence for the nucleosome’s existence.

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encounters DNA in naked form, random degradation ofthe phosphodiester backbone occurs, and a relatively ho-mogeneous distribution of DNA sizes is visualized (ifthe reaction were allowed to proceed longer, DNA wouldbe eventually degraded to mononucleotides). In contrast,when cell nuclei are treated with the same nuclease, ratherthan degrade the genome in an identical manner, the en-zyme generates populations of discretely sized DNA frag-ments that appear to occur in multiples of 180 base pairs.This suggest that in vivo, the DNA is somehow packagedinto “180 base pair installments” such that only the DNAstretch between two adjacent 180 base pair “packages” isaccessible to the nuclease. At the same time, analysis ofthe hydrodynamic properties of individual DNA–proteinparticles released by nuclease using analytical centrifuga-tions, allowed K. van Holde and coworkers to measure itsmolecular weight at ca. 180,000 Da.

Very strong support for the notion of chromatin be-ing composed of a reiteration of identical subunits camefrom electron microscopic studies by C. Woodcock andother scientists in 1973–1974. When preparations of chro-matin were spread under appropriate ionic conditions ona carbon grid and visualized under the EM, a remarkable“bead-on-a-string” fiber was visualized (Fig. 5). Cross-linking experiments by J. Thomas and R. Kornberg in-dicated that the histones’ representation in chromatin isstoichiometric; the combined weight of all these data ledto R. Kornberg’s proposal of the nucleosome hypothesis,according to which the elementary particle of chromatin(i.e., “the bead” in the electron micrograph) consisted of180 bp of DNA combined with eight molecules of corehistone and one molecule of linker histone.

D. The Structure of the Nucleosome

Data described in the preceding section set the stage for anexperimental assault on the atomic structure of the nucle-

FIGURE 5 “Beads-on-a-string”—insect chromatin visualized un-der the electron microscope (EM kindly provided by U. Scheer).

osome. Two lines of evidence obtained in the mid-1970sindicated that the histone proteins lie on the inside of thenucleosome, while the DNA is somehow exposed on itsoutside surface. M. Noll used the nuclease DNAse I todemonstrate that under appropriate experimental condi-tions, all 146 base pairs of DNA in a single nucleoso-mal particle are cleaved by this nuclease—one cleavagewas seen to occur every 10−11 base pairs. This suggestedthat the nucleic acid is exposed to solution, rather than isshielded by the histone proteins. More definitive evidenceto that effect came from biophysical studies in the labsof B. Richards and C. Crane-Robinson, who used neutronscattering to demonstrate that DNA does, indeed, lie onthe outside of the core histone particle.

In the 20 years that followed, the nucleosome was thesubject of intense investigations. X-ray crystallographicanalysis from T. Richmond and A. Klug, and subsequentlyG. Arents and E. Moudrianakis, illuminated the spatialarrangement of its constituents, as did protein-DNA cross-linking studies in the lab of A. Mirzabekov, while thedetails of the structural distortion that DNA undergoes inthe nucleosome were provided by J. Hayes and A. Wolffe.The description that follows is based on data from all ofthese studies, as well as and the highest-resolution (2.8A)X-ray crystal structure currently available (provided by T.Richmond’s research group in 1997).

1. DNA Structure in the Nucleosome

Compaction of DNA into the nucleosome involves thewinding of 146 base pairs of DNA into ca 1.7 left-handedturns around the histones (Fig. 6). Such a representationis very useful to help visualize what a nucleosome lookslike but, unfortunately, presents the erroneous view thatDNA is complacently wound onto the histones with littleor no structural stress. The reality is quite contrary to whatone could divine from this drawing: in assembling into thenucleosome, DNA is very severely distorted from its con-ventional and familiar B-form. First, to twist around thehistones, the DNA backbone has to be very severely bent—the turns that it makes likely approach the limit of thermo-dynamic feasibility. In addition, topological requirementsof winding a right-handed double helix into a left-handedsuperhelix necessitate that the DNA be partially unwoundfrom its conventional 10.5 base pairs per helix turn.

The distortion of the DNA in the nucleosome has sev-eral important functional consequences. Because DNAneeds to bend as it winds into the nucleosome, particularDNA sequences that bend more easily offer a thermody-namic advantage in this process; this means that the preciseway in which a given DNA sequence associates with thehistones—i.e., the way in which specific sequences in theDNA are rotated toward, or away from the core histones—

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FIGURE 6 (A) A schematic of the DNA path around the nucleo-some. (B) DNA distortion in one turn of the superhelix. (C) Sterichindrance in access to nucleosomal DNA.

is going to be at least in part sequence dependent. Such“rotational” positioning of the DNA relative to the core hi-stone octamer can have profound effects on the regulatorybehavior of the DNA in the nucleus, as we shall see.

Furthermore, the severe structural distortion of the DNAin the nucleosome is something of an Achilles heel, since itimplies that this entire structure is amenable to disruption.We do not wish to create the impression that the nucleo-some is intrinsically unstable—quite the contrary, an intacthistone octamer can remain complexed with DNA underconditions of physiological pH and low ionic strength forvery extended periods of time. Should the arrangementof the core histones within the nucleosome, or histone–DNA interactions themselves be altered, however, DNAwill attempt to release topological and structural stress byrecovering some B-form normalcy. This feature of the nu-cleosome is efficiently exploited in transcriptional control.

2. The Histone Octamer

The core histone octamer has a remarkably lucid struc-ture; the key to appreciating this point—possibly hidden

by the maze of barrels and ribbons in diagrams—is to real-ize that the globular domain of all core histones assumesan identical secondary and tertiary structure (Fig. 7). Inthis “histone fold” motif, the central, extended α helix isflanked on each end by a short turn (loop) of the polypep-tide, and then—again, on each end—a shorter α helix (i.e.,somewhat like the letter “Z”). The shorter helices resideon top of the same side of the longer helix—at its oppositeends, of course—and both lie at an approximately right an-gle to it. An identity of folding into this motif allows twohistone molecules to heterodimerize via a “handshake”:the central α helices of histones H3 and H4 fit togetherdiagonally to form an “X” (extending the “handshake”analogy, this is equivalent to the touching of palms), andthe shorter helices on each side of the central helix projecttoward their counterparts on the other histone molecule(analogous to fingers of one hand wrapping around theother hand). Histones H2A and H2B interact via an iden-tical handshake.

In terms of binding DNA, the most immediate result oftwo histones coming together in a structure like this one isthat the interhelical turns (loops) of each histone moleculebecome juxtaposed on each end (“top” and “bottom”) ofthe “X” that is formed. These loops build a ramp ontowhich DNA is wound (each histone dimer associates withca. 2.5 turns of DNA, i.e., ca. 27–28 base pairs); the shorterhelices on one end of each histone project toward the “frontend” of the “X” and also contact the DNA.

This histone heterodimer, then, is the elementary sub-unit of the nucleosome—the entire entity is built by fit-ting four such heterodimers together, in the followingway: two H3/H4 heterodimers come together to form theH3/H4 tetramer (i.e., an “X X” is assembled). Within thetetramer, the short α helix at the end of one histone H3molecule contacts such a helix on the other histone H3.Thus, the histones are joined end-to-end: H4′–H3′—H3–H4. The resulting entity, in fact, does occur in vivo dur-ing postreplicative chromatin assembly, when it associateswith approximately 120 bp of DNA. Its functional prop-erties, however, differ quite significantly from that of thehistone octamer bound to 146 base puirs; as shown inthe lab of J. Hansen, DNA bound to an H3/H4 tetrameris much more accessible to binding by nonhistone reg-ulators than DNA in a conventional nucleosome, andthis has immediate functional consequences in terms ofthe effects of DNA replication on genome behavior (seefollowing).

Once the H3/H4 tetramer is formed, it is joined by twodimers of H2A/H2B, primarily via hydrophobic and hy-drogen bond contacts between histone H4 with histoneH2B. The resulting daisychain of histones can be repre-sented in linear form to illuminate the pattern of histone–histone interactions (contacts made within each dimer are

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FIGURE 7 (A) The histone dimers as they interact in the nucleosome. For simplicity, only one histone set is shown.(B) The nucleosome and locations of histone–DNA interactions.

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represented by a shorter dash, and between dimers—by alonger one):


In vivo, of course, the histones are not arranged in linearfashion, but rather bundle together into a rather compactparticle: the H3/H4 tetramer forms an inverted V (i.e., acapital lambda, �), and the H2A/H2B heterodimers at-tach to either side of the cleft at the bottom of this “�.”Viewed from the side, the resulting entity is shaped like awedge (the octamer tapers toward the side of the H3/H4tetramer). Yet another unfortunate consequence of text-book schematics is that the octamer is commonly repre-sented as a hockey puck—i.e., somewhat of a monolithicentity. The octamer is not a monolith—in fact, it is veryclear that the heterodimerization interface between his-tone H4 and histone H2B is a somewhat delicate one,and is amenable to disruption. Thus, a better analogy forthe octamer would be that of a Rubik’s cube—a three-dimensional jigsaw puzzle—with individual componentsfitting together in a dynamic, pliable arrangement.

Sixteen loop motifs decorate the sides of the octamerside like rungs on a ladder, and these offer an interactioninterface to the DNA. The majority of contacts between thehistones and the DNA occur via hydrogen bonds and saltlinks to the phosphates in the DNA backbone, althoughtwo additional important sets of interactions occur whenthe side chain of arginine in the histone penetrates the mi-nor groove of the DNA, and when, remarkably, nonpolarcontacts are made with the deoxyribose.

FIGURE 8 The histone tails shown schematically to their full predicted length in relation to the nucleosome coreparticle (drawn to scale).

From a regulatory perspective, as shown in Fig. 6, a dra-matic consequence of DNA winding onto the octamer is achange in its accessibility to solution (and, by extension,to nonhistone protein regulators). The obstacles are manyin nature: the octamer itself obstructs the face of the DNAthat is turned toward it, and—because of the proximity ofthe DNA gyres in the superhelix, one turn of the DNAimpedes access to the adjacent DNA turn. A great dealof experimental attention has been directed, therefore, to-ward understanding how particular sequences bind to thehistone octamer, and how specific regulators then bind tonucleosomes containing their target sites.

One important experimental variable in these assayscannot, unfortunately, be seen in the X-ray crystal struc-ture: the NH2-terminal core histone tails account for ca.25% of the histone mass, but are mostly unstructured inthe crystals. It is known that the tails of histones H2B andH3 do not stick out to the side of the nucleosome, butrather emerge into solution between the gyres of the DNAsuperhelix. The whereabouts of the histone H4 tail is un-clear (with the exception of a small segment that makesan internucleosomal contact in the crystal!).

Attention to the tails’ structure is justified by theirprominence in regulatory phenomena that occur on chro-matin (Section IV.A). While normal nucleosome assemblycan occur on tailless (proteolyzed) histones, normal tran-scriptional control can not—and the regulatory reach ofthe tails can be illustrated by their sheer stereochemicalreach (Fig. 8). Because the tails contain a combined 44 ly-sine residues, at physiological pH they very likely coat the

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underlying DNA like a maze of positively charged tenta-cles. Thus, the alterations in the charge of the tails effectedby postranslational covalent modifications are likely tohave impact on chromatin.

E. “. . . Not by Beads Alone”: Higher-OrderChromatin Structure

It is clear that the nucleosome as a tool is not sufficient tocompact the entirety of the genome into the nucleus; quitea feat of condensing is required to convert the “beads-on-a-string” fiber into the dramatic metaphase chromosomesso familiar to many (if the entirety of the genome wereassembled into a fully extended nucleosomal array, itslength would be ca. 15 cm).

One important functional component of further foldingby the nucleosomal fiber is the linker histone (H1). Asits name implies, its binding site is with the DNA stretchbetween two adjacent octamer particles; the precise man-ner in which the linker histone binds to that DNA hasbeen the subject of much investigation and controversy(Fig. 9). Whatever the precise mode of its associationwith DNA, one important consequence of this associationis charge neutralization over the linker DNA stretch—thatis, the shielding of the negatively charged phosphate back-bone from solution. Experiments in the laboratory of M.Gorovsky have shown that the ciliate Tetrahymena can sur-vive without linker histone, but that the size of its nucleusincreases twofold; furthermore, mutations that increasenegative charge on histone H1 (mimicking its phosphory-lation) promote its loss from chromatin. In addition, dataon the folding of chromatin in vitro in the presence andabsence of linker histone are also fully consistent withthe model that the presence of histone H1 is required forproper folding of the “bead-on-a-string” fiber.

What exactly this fiber folds into is—remarkably—notknown. A great variety of EM and other approaches—dutifully represented in textbook schematics—have sug-gested that the next level of chromatin compaction is anentity termed “the 30-nm fiber.” In 1979, F. Thoma, A.Koller, and A. Klug proposed a model for this entity ac-

FIGURE 9 Two proposed models for the location of the linkerhistone (black sphere) relative to the nucleosome core particle.

FIGURE 10 Model for higher-order chromatin folding.

cording to which the nucleosomal fiber winds onto itselfto form a solenoidal structure, with ca. six nucleosomesper turn of the superhelix. Remarkably, whether such anentity forms in vivo, and what the precise arrangement isof the nucleosome within this structure, remains an openissue, and other models for this fiber have been proposedby C. Woodcock and several other scientists.

Beyond the mysterious 30-nm fiber lies an undiscoveredcountry of dramatic proportions—we currently lack thetechnical tools to examine higher-order chromatin fold-ing. It has been proposed (Fig. 10) that large domainsof chromatin emerge from a central scaffold in the formof loops, and work from the labs of U. Laemmli and J.Sedat presented evidence in support of this model. Detailsremain very elusive, however. It is important to realize,however, that even in the absence of information abouthigher-order chromatin structure, our current understand-ing of the nucleosome and the nucleosomal fiber offers anample stage for the unfolding of very complex gene reg-ulatory phenomena concomitant with chromatin structuretransitions. These are reviewed in the next section.


It is somewhat ironic that the discovery of the nucleosome,and the determination by M. Noll in 1974 of its ubiquityin the genome, were partly responsible for the transientelimination of histones from the stage on which transcrip-tional control was thought to unfold. In retrospect, it iseasy to see why: because all nucleosomes are the same(in a certain sense, they are), it was very hard to imagine,

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how anything pertinent to gene-specific regulation couldoccur on such a homogeneous, monotonously reiterativesubstrate as the “beads-on-a-string” fiber. Thus, all tran-scriptional and other regulatory phenomena were thoughtto occur on nucleosome-free stretches of “naked” DNA,although it remained unclear, whether such stretches canbe found in vivo.

This notion was also supported by experimental ob-servations on nonhistone factor interactions with chro-matin templates in vitro. Such analysis indicated that manyproteins—for example, the TATA-box binding protein,TBP—cannot bind to their DNA sites when they are as-sembled into nucleosomes. From these, and other obser-vations, emerged a model according to which inactive re-gions of the genome were packaged into nucleosomes;chromatin, thus, was viewed as a general (nonspecific),repressive entity. The elimination of histones from activeregions of the genome was then thought to set the stagefor binding by nonhistone factors and transcriptional ac-tivation via the recruitment of RNA polymerase.

Over the past years, however, several lines of evidenceemerged that suggested this model was an oversimplifi-cation. It became apparent that chromatin structure is notmonotonous or homogeneously isomorphous; instead, ex-tensive localized alterations in its nucleoprotein fiber wereobserved concomitant with changes in genomic activityat specific loci. In an important complementary develop-ment, a host of macromolecular complexes were discov-ered that populate the nucleus and effect these alterations.In addition, specific gene loci were shown to have propertranscriptional control depend on their assembly into chro-matin. Finally, whole-genome expression profiling experi-ments described above provided strong evidence that chro-matin is not a generalized repressor, and that many genesrequire chromatin for proper regulation.

A. Chromatin Structure AlterationsThat Occur in vivo

While structural studies of the nucleosome provided veryimportant information for scholars of transcription, a ma-jor issue for understanding how the genome behaves inchromatin form was to learn what—if anything—happensto chromatin during gene activation and repression in vivo.The cytological studies described earlier in this article(Section II) indicated that localized alterations do occur,but the resolution limitations inherent in such methodsprevented an interpretation of the data in molecular terms:for example, puffs that form on polytene chromosomesare very conspicuous, but what exactly happens to thechromatin fiber during puffing remained unclear (and, in-cidentally, still does).

1. Remodeling

As was the case with the initial identification of subunitcomposition of chromatin, the first clues to chromatin re-modeling phenomena concomitant with alterations in geneactivity came from the use of nucleases. In the late 1970s,C. Wu and S. Elgin, and, independently, S. Nedospasovand G. Georgiev used nucleases such as DNAse I and mi-crococcal nuclease (MNase) to probe chromatin in vivo.The notion behind this approach was the use of low quan-tities of enzyme—the prediction being that mature chro-matin, i.e., DNA that is tightly complexed with histones,would not be accessible to cleavage by nuclease, and thusonly DNA stretches that were not assembled into conven-tional chromatin would be “visible” to the nuclease. Theseinvestigators used an elegant technical trick—“indirectend-labeling”—to reveal such DNA stretches. Wu andNedospasov made the same observation—that regulatoryDNA (for example, promoters of active genes, or of genesthat are poised for upregulation) is preferentially accessi-ble to such nucleases and forms a “nuclease hypersensitivesite” in vivo. At the time, the structural basis of this en-tity was unclear; remarkably, it continues to be not entirelycertain to this day, although we have a much deeper under-standing of the molecular machines responsible for effect-ing this chromatin structure alteration and of the propertiesthese machines exhibit in vitro.

An appreciation for the role of such hypersensitive sitesto gene regulation grew in the early 1980s, when it wasdiscovered that they are a relatively ubiquitous featureof promoters and enhancers in the eukaryotic genome.A seminal observation was made in 1984 using a modelsystem that has provided many insights into the role ofchromatin in gene control: the regulation of transcriptionof the mouse mammary tumor virus (MMTV) genome bythe glucocorticoid receptor (GR) and its ligands, the gluco-corticoids (cortisol, corticosterone, and aldosterone). GRis a small-molecule-regulated transcription factor: inac-tive in the absence of hormone, it translocates to the nu-cleus in its presence, binds to target genes, and activatestranscription. In 1974, it was discovered that transcriptiondriven by MMTV—the etiologic agent of breast cancerin mice—is rapidly upregulated by treatment with a syn-thetic GR ligand, dexamethasone. Ten years later, K. Zaretand K. Yamamoto discovered that hormonal treatment in-duces a strong DNAse I hypersensitive site in the MMTVpromoter (called the “long terminal repeat,” or LTR), andthat this hypersensitive site vanished rapidly upon removalof hormone. This established a correlation between theextent of such remodeling and the level of transcriptionat this locus. An example of a DNAse I hypersensitivesite induced by a nuclear hormone receptor is shown inFig. 11.

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FIGURE 11 Binding of the thyroid hormone receptor (TR) tochromatin generates a DNAse I hypersensitive site (arrowheads).“TRE” represents the TR binding site.

It is important to note that the disruption of chromatinstructure that is described as a “hypersensitive” site is ahighly localized event and only affects ca. 50–500 basepairs of DNA (i.e., only several nuclesomes’ worth)—thus, while the insect relative of GR, the ecdysone recep-tor, functions via highly similar mechanisms, it should notbe inferred that polytene chromosome puffing observedsome 25 years prior by Clever and Karlson in Chironomusafter ecdysone injection (see Section II) is due to suchremodeling—puffing is a large-scale event which affectsstretches of the genome that are many thousands of bplong. A molecular correlate to “puffing” was discoveredin the lab of C. Crane-Robinson in 1994 in studies ofthe globin gene cluster. A marvel of gene regulation, theglobin genes in higher vertebrates have been investigatedin great detail, in part because they represent one of the bestcharacterized instances of a “chromosomal domain”—alarge continuous stretch of the genome in which severalgenes reside whose expression is coordinately regulated(for instance, in mammals, a progression of gene activationoccurs, with “fetal” globin genes active during embryonicdevelopment, and a switch to “adult” type globin genesafter birth). In the chicken genome, the β-globin locusencompasses ca. 33,000 base pairs; by using nucleases,Crane-Robinson’s research group showed that in erythro-cytes (i.e., the cell type that transcribes globin genes) thisentire portion of the chromosome was more sensitive tonucleases than a transcriptionally inert one.

We emphasize that the distinction between nucleasesensitivity and hypesensitivity is not merely a semanticone. During the activation of gene expression in vivo, itis quite likely that as a first step, a large stretch of chro-matin undergoes some structural transition to a more ac-cessible conformation (it is possible that such a transi-

tion is mechanistically similar to that between heterochro-matin and euchromatin). Within such a large domain ofless compacted chromatin, targeted chromatin remodelingover short DNA segments then induces hypersensitivity,i.e., a more dramatic disruption of histone–DNA contacts.

2. Modification

An additional important observation on the β-globin locusrelated to a different type of chromatin structure alterationobserved in vivo: a change in the covalent modificationstatus of histone tails within a domain of transcription-ally active chromatin. To better illuminate the significanceof this finding, we must briefly review the biochemistryof such modifications. As mentioned earlier, in 1964 itwas discovered that particular lysine residues in the NH2-terminal tails of the core histones are reversibly covalentlymodified by acetylation:

· · · NH+3 + Ac-CoA · · · NH2 CO CH3.

This observation instantly suggested an electrostaticmechanism for gene regulation: the histone tails carry awealth of positive charge on their basic amino acids andare thus expected to bind tightly to the negatively chargedphosphates in the DNA backbone. Acetylation eliminatesthe charge on the target amino acid side chain, and thusa hyperacetylated histone tail would be expected to bindless tightly to the DNA, and make it more visible to theintranuclear world. Thus, there must be a correlation be-tween chromatin hyperacetylation and transcriptional acti-vation (and, conversely, between chromatin deacetylationand transcriptional repression).

This model makes a lot of sense from first principle apriori considerations, but is it supported by experimentalevidence? The correlation between levels of acetylationand transcription clearly exists in vivo. A very powerfultool in making this experimental determination has beenantibodies that selectively recognize hyperacetylated ordeacetylated histone tails. These have been used with greatsuccess in two experimental strategies. The first, fluores-cent in situ hybridization (FISH), is cytological: it allowsone to examine histone acetylation status over entire chro-mosomes. In the most general sense, it involves the prepa-ration of nuclei from cells (or tissue) under biochemicallymild conditions, the immobilization of their chromatincontent on a suitable support, and the probing of the re-sulting karyotype with an antibody of choice, followed byprobing with a “secondary” antibody that reveals wherethe original antibody bound. The secondary antibody ischemically coupled to a fluorescent dye, and this allowsvisualization under a microscope of entire chromosomeswith brightly fluorescent segments that have bound to theantibody.

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822 Chromatin Structure and Modification

A model system best suited for such analysis is onewhere an entire chromosome has an altered expressionstate; the best characterized example comes from stud-ies of dosage compensation: this evolutionarily con-served device for coping with a different autosome to sexchromosome ratio between genders in metazoa changesexpression levels of the X chromosome depending on gen-der. For example, in mammals, one of the two X chromo-somes in females is inactivated (thus, identical expressionlevels are achieved for X-linked loci between males andfemales). In insects, on the other hand, the single X inmales is transcriptionally upregulated twofold, thus ad-justing its expression level to that of two X chromosomesin females. FISH analysis by B. Turner and colleaguesdemonstrated that the inactive X in mammalian femalesis hypoacetylated; this provides an important correlate toOhno’s and Lyon’s observations (v.s.) from the 1960s thatthis chromosome is condensed into heterochromatin andtranscriptionally silenced. On the other hand, FISH anal-ysis in Drosophila revealed that the transcriptionally “hy-peractive” X chromosome in males is hyperacetylated rel-ative to X chromosomes in females.

The second experimental approach that revealed acorrelation between states of acetylation and levels oftranscriptional activity is chromatin immunoprecipitation(ChIP). This method was developed by M. Solomon andA. Varshavsky, and further by D. Allis and M. Gorovsky.This technique allows one to detemine if a protein of in-terest interacts in vivo with a DNA stretch of interest; thereagents required for such analysis are an antibody againsta particular protein and knowledge of the primary DNA se-quence of the locus of interest. This ingenious and power-ful method begins by taking an in vivo snapshot of protein–DNA and protein–protein interactions in the nucleus viaa brief incubation of living cells or tissue in formalde-hyde; this small molecule rapidly penetrates into the celland introduces covalent crosslinks between proteins andDNA that they are bound to, as well as between proteinsthat are in sufficient physical proximity (i.e., are in a com-plex). Chromatin is then isolated from cells and sheared(by acoustical means, i.e., sonication) into small—ca. 500base pairs—fragments. An immunoprecipitation is thenperformed to isolate from this complex mixture the pro-tein of interest (and whatever happens to be covalently at-tached to it): the antibody is immobilized on a suspensionof agarose beads, and the beads are then mixed extensivelywith the sonicated chromatin to allow the antibody to bindits antigen. The beads are then isolated by centrifugation,which redistributes the target protein from solution into thepellet (together with the beads). The cross links betweenthe protein and DNA are then eliminated, and the DNAisolated. Finally, the presence of a given DNA sequencein this isolate is assayed by PCR or by Southern blotting.

ChIP has been applied extensively to analyze the hi-stone tail acetylation status over particular stretches ofvarious genomes (from budding yeast to humans). Thegeneral conclusion from these experiments is that, indeed,transcriptional repression is accompanied by localized hi-stone deacetylation, while transcriptional activation oc-curs within loci that are associated with hyperacetylatedhistones: for example, theDNAse I sensitive domain ofthe active chicken β-globin locus that was described ear-lier was found to be hyperacetylated by ChIP analysis (C.Crane-Robinson and colleagues), while transcriptionallysilent stretches of yeast chromatin, such as the mating typeloci, are deacetylated (J. Broach and colleagues).

It is quite striking that our current biochemical insightinto the enzymatic reaction of histone tail acetylation,the causative agents of this modification (Section IV.C),and their involvement in transcriptional control in vivo isnot paralleled by a similar understanding of the structuraleffects of histone tail hyperacetylation on chromatin, or ofthe mechanistic underpinnings of the general stimulatoryeffect that this modification has on the transcriptionalmachinery.

The structural puzzles come in part from the fact thatthe tails fail to appear in X-ray crystallographic analysis.It is clear that in the case of histones H3 and H2B, thetails emerge into solution by passing through the twoadjacent DNA double helices that lie on the surface of theoctamer, but their subsequent path—assuming a definedone exists, which is not at all clear—is unknown. Inaddition, a short segment of the histone H4 tail—sevenamino acids—can be seen making contact with a histoneH2A/H2B dimer in an adjacent nucleosomal particle.How—or whether—the tails engage the DNA of thenucleosome they belong to is unknown.

Faced with a void of structural understanding from crys-tallographic analysis, scientists turned to other biophysi-cal methods to investigate what happens to the tails uponacetylation. In 1982, E. M. Bradbury and colleagues usedNMR analysis of peptides corresponding to the histoneH4 tail, and found that it bound only weakly to DNA, andthat hyperacetylation abolished this binding. By thermaldenaturation analysis these scientists derived a quantita-tive estimate of the effect of acetylation: the intact pep-tide was seen binding to DNA with an affinity of 50 pM,while acetylation reduced it to 10 µM! The magnitudeof this effect was subsequently shown to depend on thenumber of lysine residues acetylated, and in the contextof the nucleosome—rather than as isolated peptides—thehistone tails continued to make some contacts with theDNA even when hyperacetylated. In studies using circulardichroism spectra (J. Parello and colleagues), DNA-boundstretches of both histone H4 and H3 tails were found tobe highly structured and adopt an α-helical conformation,

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while those of histones H2A and H2B were seen to as-sume a random coil conformation. The working modelas to the local effects of acetylation on tail structure andinteraction with DNA, then, suggests that it may disruptthe (currently unknown) secondary structure the tails as-sume, and lessen the extent to which the tails interact withDNA.

What is the relevance of these observations to transcrip-tional control? If, indeed, the tails become more looselyassociated with DNA upon hyperacetylation, it is possi-ble that the underlying DNA becomes more accessible tononhistone regulators. In vitro experiments with purifiedchromatin components and particular transcriptional reg-ulators (A. Wolffe, J. Workman, and their colleagues) havefound that histone hyperacetylation potentiates binding tonucleosomal substrates by such proteins as TFIIIA, Gal4,and USF. Whether such potentiation of binding occurs invivo is unknown.

As mentioned earlier, a segment of the histone H4 tailwas seen making an internucleosomal contact in the crys-tal structure. These data, and other observations, lent sup-port to the hypothesis that tail acetylation affects higher-order folding of chromatin. Strong evidence to this effectwas obtained by J. Hansen and his colleagues, who usedanalytical ultracentrifugation to demonstrate that the hy-drodynamic properties of chromatin fibers can be mod-ulated in vitro. An alteration in the ionic strength of thesolution dramatically altered the shape of the nucleosomalfiber: as the concentration of Mg2+ increased, the fiber be-came much more compact. Importantly, the deacetylationof this fiber was then shown to have an identical effect:thus, hyperacetylated chromatin adopts an extended con-formation, and deacetylation promotes folding. It is possi-ble, therefore, that the increased accessibility to nucleasesseen in transcriptionally active, hyperacetylated chromo-somal domains in vivo is causally linked to a change inthe extent of chromatin compaction that can be observedin vitro. Additional support for such a connection comesfrom the work of A. Belmont and coworkers, who usedin vivo imaging techniques to demonstrate that transcrip-tional activation is accompanied by the hyperacetylationand unfolding of a chromosomal domain spanning severalhundred thousand base pairs. Most remarkably, this pro-cess occurred even in the absence of transcription—thisimportant control experiment indicated that the unfoldingis not a consequence of the passage by the RNA poly-merase II complex (an imposing entity) through the chro-mosome, and is an independently regulated phenomenon.

Whatever the structural consequences of acetylation onchromatin, some of the strongest evidence in favor of itsdirect role in controlling the genome comes from data il-luminating the abundance inside the nucleus of enzymeseffecting histone tail modification, and from experiments

that show these enzymes to be directly involved in tran-scriptional regulation in vivo. These are reviewed in thenext section.

B. Chromatin Disruption and Modification:The Enzymatic Machinery

In interesting testimony to the powerful influence ofmethodology over the course scientific inquiry, investiga-tions of the molecular agents that effect chromatin struc-ture alterations in vivo have followed a remarkably uni-form scheme, somewhat reminiscent of a Bildungsroman.It begins with a genetic screen in budding yeast for strainsthat exhibit particular phenotypes related to gene control(e.g., the ability to activate a particular metabolical path-way), the molecular cloning of the underlying loci, andtheir subsequent putative identification as transcriptionalregulators. Next, analysis in metazoa reveals the existenceof homologs, and an enzymatic assay is developed thatdemonstrates that these proteins have the capacity to alterchromatin structure. Mutational analysis reveals that thereis a correlation between the enzymatic activity of the pro-tein and its ability to act in transcriptional control. Bio-chemical analysis of whole-cell extracts then finds theseproteins within large, multisubunit complexes, only a fewpolypeptides in which have enzymatic activity, while therest are of uncertain function. Finally, in vitro experimentsare done that show these complexes can be targeted byparticular transcriptional activators or repressors, and thatmutations in these transcription factors that alter the abil-ity to interact with chromatin modifying and remodellingcomplexes impair their properties as regulators.

1. Disrupt and Conquer: ATP-DependentChromatin Remodeling Engines

In the 1980s, genetic studies in laboratories of M. Carlsonand F. Winston identified a number of mutant yeast strainsthat failed to metabolize sucrose; following convention,the many loci revealed in the screen were called SNF(sucrose nonfermenter) and given a number (i.e., SNF2,SNF5, etc.). At approximately the same time, the lab of K.Nasmyth discovered a number of loci in the yeast genomethat when mutated, incapacitated the switching of mat-ing type; these were dubbed SWI (for “switch”) and alsonumbered (SWI5, SWI2, etc.). The capacity to metabolizesucrose is dependent on the activation of specific genesthat belong to the cognate metabolic pathway (in partic-ular, an invertase); the switching of mating type requiresthe upregulation of a gene that encodes an endonucle-ase required for initiating a DNA recombination reaction.Thus, it appeared that a common feature of the two other-wise quite unrelated phenotypes in the mutant strains (fail-ure to metabolize sugar or the incapacity to mate) was a

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deficiency in gene activation; most remarkably, the samelocus was commonly found mutated in both screens. Thecognate gene—SWI2/SNF2—was subsequently found inwork from I. Herskowitz’s laboratory to be required fortranscriptional upregulation of a number of budding yeastgenes.

In a landmark set of experiments, the lab of F. Winston in1992 demonstrated that the action of SWI2/SNF2 in geneactivation is mediated via alleviating the repressive effecton gene expression of chromatin structure. These studiesused the classical genetic notion of epistasis—i.e., the ca-pacity of mutations in one locus to mask a mutation in adifferent locus—to uncover genetic interactions betweenSWI2/SNF2 and the genes for histones: it was shown thatthe inability of certain genes in the yeast genome to beactivated when SWI2/SNF2 is mutant can be “healed” bymaking mutations in histone genes (for example, by low-ering the histone content of the nucleus). Most strikingly,subsequent analysis by I. Herskowitz, C. Peterson, and M.Osley demonstrated that even less drastic measures—forexample, point mutations in histone H4 that impair the ca-pacity of H2A–H2B to bind the (H3/H4) tetramer—willalso make SWI2/SNF2 unnecessary. Thus, those genes inyeast that require SWI2/SNF2 to become transcriptionallyactive lose that requirement when chromatin structure isdestabilized by making mutations in histone genes.

The most immediate prediction of these remarkableexperiments—consider the extraordinary fact that the en-tirety of chromatin within the yeast nucleus can be alteredby genetic means—is that the product of the SWI2/SNF2gene somehow alters chromatin structure over target genepromoters. A great number of studies have yielded datafully compatible with that notion (the laboratories are toomany to list, but, in addition to those already mentionedinclude those of G. Crabtree, W. Horz, R. Kingston, C.Peterson, J. Workman).

Several important facts emerged regarding SWI2/SNF2.Biochemically, it was found to be an ATPase (i.e., it hy-drolyses ribo-ATP to release ADP and phosphate)—thiswas an important observation, because it illuminated apossible requirement for energy in chromatin remodeling.In vivo, it was discovered as one of the core componentsof a multisubunit complex designated as SWI/SNF (pro-nounced “switch-sniff”). In vitro assays with nucleosomaltemplates and transcriptional regulators demonstrated thatSWI/SNF can remodel histone–DNA contacts such thatsubsequent access by these regulators to the remodelednucleosomal template is increased. For example, a TATAbox buried within a nucleosome became more accessi-ble to TBP after transcriptional activator-dependent ac-tion on that nucleosome. The exact structural nature of the“remodeled” nucleosome is unclear; it is known that thehistone–DNA contacts are loosened sufficiently to allow

access and cleavage by DNAse I, but it is also clear that thehistones remain in some contact with the DNA. Whateverthe nature of the remodeled entity, energy derived fromthe ATP hydrolysis is required for its generation.

What relevance does such action of SWI/SNF in vitrohave to transcriptional control in vivo? The best explana-tion we can offer comes from studies in mammalian sys-tems. Thus, work by G. Hager and colleagues on transcrip-tional control of the MMTV promoter (Section IV.A.1)focused on the function of the DNAse I hypersensitivesite that is induced by the liganded glucocorticoid receptorconcomitant with transcriptional activation. An importantclue came from a comparative analysis of MMTV tran-scription on copies of the viral genome that have beenintegrated into the chromosome (and thus assume nativechromatin organization) vs. such DNA that has transientlyintroduced into the cell by a technique called transfection(the DNA remains extrachromosomal and does not assem-ble physiological chromatin; it is lost from the cell afteronly a few rounds of cell division). It was known thatfull-scale activated transcription on the MMTV promoterrequired an activator called NF1; remarkably, NF1 actionwas only dependent on GR and its ligand when the targetpromoter was chromatinized: on transiently transfectedDNA, NF1 was able to bind to DNA without any abettingaction from the receptor.

An explanation for this interesting synergy was pro-vided in a famous study by T. Archer and G. Hager in 1992:they proposed that the MMTV promoter adopts a nonran-dom chromatin organization in vivo, such that the bindingsite of NF1 is occluded by a nucleosome. In contrast toNF1, GR can directly bind to chromatin over the MMTVpromoter, and then somehow remodels histone–DNA con-tacts adjacent to its binding sites. This remodeling (man-ifested as a DNAse I hypersensitive site) facilitates NF1access and potentiates transcriptional activation (Fig. 12).This two-step (“bimodal”) mechanism for GR action ledto several predictions: the receptor had to be shown ascompetent for binding to nucleosomes, and also for therecruitment of a chromatin remodeling engine. In fact,GR and several other members of the nuclear hormone re-ceptor superfamily can bind to nucleosomes in vitro—thisis quite an achievement, considering the extensive sterichindrance exerted by the nucleosome ( an example of suchbinding to nucleosomes by the thyroid hormone receptoris shown in Fig. 13). In addition, T. Archer and colleaguesdemonstrated that a human homolog of the budding yeastSWI/SNF complex is required for transcriptional activa-tion of MMTV by GR. A central conclusion of this analy-sis is that bona fide transcriptional control of this promotercannot be recapitulated on naked DNA, and that the infras-tructure of chromatin is integrated into the transcriptionalregulatory pathways affecting MMTV.

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FIGURE 12 A schematic of the low-resolution map of the MMTV promoter (LTR); the position of the transcriptionstart site (“+1”), the GR binding site, the NF1 binding site, and the histone octamers are shown.

Whether SWI/SNF is directly responsible for the chro-matin remodeling over this, or any other mammalian pro-moter in vivo remains an open issue. Work from O. Delat-tre and colleagues showed that truncating mutations in acore subunit of the human SWI/SNF complex cause a dev-astating disorder—“malignant rhabdoid tumors” (MRT);these develop on the kidneys and in the central nervoussystem in infants (onset occurs before 2 years of age) andare highly aggressive. This indicates that the maintenanceof normal cell proliferation and differentiation status inhumans require the function of the SWI/SNF complex.It is not clear, however, if the facilitation of access viathe remodeling of histone–DNA contacts accounts for theentirety of action by SWI/SNF during activation.

For specific promoters in budding yeast, it has beenshown that ablation of the SWI/SNF complex causes thesepromoters to become inactivated due to a failure to re-model chromatin: no DNAse I hypersensitive site is visual-ized over these promoters in the absence of SWI/SNF. Onemysterious issue concerning the function of SWI/SNF isits role in transcriptional repression; whole-genome ex-pression analysis in the laboratory of R. Young demon-

FIGURE 13 The thyroid hormone receptor can bind to naked DNA containing its response element (left half), andalso to this DNA when it is assembled into a nucleosome (right half).

strated that a number of budding yeast genes becometranscriptionally activated in strains lacking SWI2 (anda different set of genes become silenced). While thephenomenon is clear, its mechanistic underpinnings arenot. Elegant genetic analysis in F. Winston’s laboratorydemonstrated that SWI/SNF acts directly on those pro-moters that are upregulated in its absence (i.e., it is a directrepressor of those genes, and not, for example, an activatorof a repressor). It is possible that the repressive action ofSWI/SNF has to do with its ability to effect nucleosomemobilization: the sliding of the histone octamer in cis rel-ative to the DNA. Thus, perhaps, SWI/SNF actively repo-sitions nucleosomes over particular gene promoters suchthat important regulatory DNA stretches are occluded.

Concluding our overview of ATP-dependent chromatinremodeling machines, we note that eukaryotic genomesare populated with relatives of the SWI2/SNF2 ATPases(the best characterized such relative is the protein ISWI—pronounced “eye switch”). These also occur in a large,multisubunit complexes, but their role is unclear. Many ofthese complexes have the interesting ability to organizechromatin—i.e., introduce proper spacing into disordered

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FIGURE 14 The chemical equation of ε-acetyllysine synthesis and degradation by HATs and HDACs.

nucleosomal arrays—but what these complexes actuallydo to the chromatin fiber in vivo remains to be investigated.

2. The Power of Amendments: HATs and HDACs

The extensive evidence connecting histone tail acetylationwith transcription (Section IV.A.2) suffered from its inher-ently correlative nature—it was not clear, whether changesin acetylation status were causative to, or, instead, an ef-fect of, changes in gene activity. This situation changedquite dramatically in 1996.

As shown in Fig. 14, the reaction of acetylation is cat-alyzed by histone acetyltransferases (HATs) and reversedby histone deacetylases (HDACs). In the cases of bothclasses of proteins, their enzymatic activity was used as atool to purify them from cells. The laboratory of C. D. Allisdeveloped a very unusual method for the detection of HATactivity: the in-gel assay. In this technique, a conventionalpolyacrylamide gel is prepared with a large quantity ofhistone protein polymerized directly into the gel matrix!A crude protein mixture is then resolved on the gel, andthen an in loco reaction is performed by bathing the gelin 3H-acetyl-CoA: the assumption is that the presumptiveHAT will, after having migrated to a defined position inthe gel, utilize the histone substrate all around it in the gel,and label it. The position of the HAT activity, therefore,can be visualized by fluorography. Allis’s lab identifieda major source of HAT activity in nuclei of the ciliatedprotozoan Tetrahymena that migrated at 55 kDA; its pu-rification and primary sequence characterization yieldedthe striking observation of its high homology to a knownyeast protein, Gcn5p.

To appreciate the significance of this discovery, it ishelpful to briefly review what was then known aboutGCN5 (in budding yeast, gene loci are designated by up-percase italics, i.e., ABC1, while the protein product ofthat locus is written as “Abc1p”). This gene was isolated

in the mid-1980s in a screen for strains that would havedeficiencies in regulating enzymes involved in amino acidbiosynthesis; nothing was known about the mechanismof its action until the lab of L. Guarente demonstratedin 1992–1994 that it is involved in transcriptional controlby such transcription factors as Gal4p and Gcn4p. Thegenetic approach that led to this discovery is very infor-mative: the chimeric transcriptional activator Gal4-VP16carries the DNA binding domain of the yeast transcriptionfactor Gal4p, and the transcription activation domain ofa protein from herpes simplex virus (HSV)—virally de-rived proteins and regulatory DNA stretches have evolvedto evoke large-scale responses from the transcriptional ma-chinery, and thus are very commonly used in biology. Anunexpected property of Gal4-VP16 was its toxicity to theyeast cell; work from M. Ptashne’s lab gave rise to the no-tion of “squelching”—competition by a given transcrip-tion factor for some limiting general component of thebasal transcriptional machinery. Thus, it was thought thatthte VP16 domain “squelches” important regulatory pro-teins away from other yeast promoters. Thus, reasonedGuarente and his colleagues, whatever protein—whenoverexpressed—suppresses Gal4-VP16 toxicity must berelevant to transcriptional control. Indeed, such a screenyielded the GCN5 locus; it was suggested that Gcn5p,therefore, is a “transcriptional adaptor”—an entity thattransduces the regulatory signal being sent by the DNA-bound regulator to the basal transcription machinery.

Thus, the discovery of Allis and coworkers shone in anew light: a protein found in ciliates enzymatically capa-ble of hyperacetylating histones has a very close relativein budding yeast—a protein that is required for transcrip-tional activation and is thought to be an adaptor! This wasthe first instance of a direct connection between a spe-cific HAT and transcriptional control. In subsequent years,Gcn5p in yeast and other organisms has been subjected toan extraordinarily comprehensive set of studies that firmly

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established its HAT activity as essential to transcriptionalactivation. For example, work from Allis and coworkersshowed that GCN5 is required for the transcriptional up-regulation of a number of budding yeast genes, and thatpoint mutations in Gcn5p that abrogate its catalytic activ-ity as a HAT abolish its capacity to abet transcriptionalcontrol. In addition, ChIP analysis demonstrated that his-tones over promoters of genes that are targets for bind-ing by Gcn5p-recruiting activators are hyperacetylatedconcomitant with its action. Finally, the laboratory of S.Roth reported an experiment that is the scientific equiv-alent of a coup de grace: a yeast strain was engineeredin which specific lysines in histone tails were mutatedto a noncharged residue—thus, these cells had “geneti-cally hyperacetylated histones.” In such a strain, formerlyGCN5-dependent genes lost their requirement for Gcn5pto become transcriptionally active!—thus, it was formallyproven that the in vivo function of Gcn5p at target genepromoters is to hyperacetylate the histones.

While original analysis revealed a single major HAT inTetrahymena extracts, subsequent work revealed the factthat eukaryotic genomes contain a large number of pro-teins possessing HAT activity. Importantly, an overwhelm-ing majority of these can interact with various transcrip-tion activators (hence the term “coactivator” that is used todescribe the HATs). In metazoa, the most-studied HAT isthe global transcriptional coactivator p300 and the closelyrelated protein CBP; this large protein (ca. 2400 aminoacids) contains at least four distinct interaction interfacesthat allow it to be targeted by an extraordinary varietyof transcriptional regulators, including proteins that reg-ulate the cell cycle (such as c-jun and c-Fos), cell differ-entiation (MyoD), cell-cycle checkpoints (p53), and thenuclear hormone receptors. Most importantly, as discov-ered in the laboratory of Y. Nakatani in 1996, from anenzymatic standpoint CBP is a HAT capable of hyper-acetylating all four core histones in solution. The ubiquityof CBP/p300’s involvement in transcriptional regulatorypathways in vivo is powerful evidence to the pervasive useof targeted chromatin remodeling to effect gene control. Inhumans, mutations in the gene for CBP cause Rubinstein–Taybi syndrome—a multisymptomatic disorder character-ized by mental retardation and a complex pattern of pro-found developmental abnormalities.

A well-characterized group of transcription factors thatuse HAT targeting to effect transcriptional activation arethe nuclear hormone receptors—these recruit such HATsas SRC-1 and ACTR. Thus, HATs are an integral compo-nent of signal transduction pathways involving major reg-ulators of mammalian physiology—glucose metabolism(glucocorticoids), ovulation (progesterone), developmentof secondary sexual characteristics (estradiol and testos-terone), bone morphogenesis (vitamin D), brain develop-

ment and skeletal maturation (thyroid hormone), and oth-ers. Point mutations in the gene for thyroid hormone recep-tor (TR) lead to the clinical disorder of resistance to thyroidhormone—patients present with goiter, stunted growth,and attention deficit hyperactivity disorder. Biochemicalanalysis in V. K. Chatterjee’s lab revealed that the muta-tions impair the capacity of TR to recruit HAT coactivatorsand lower—or abolish, in some cases—its ability to acti-vate transcription. These observations present very strongevidence that HAT targeting by specific transcriptional ac-tivators occurs in vivo and is relevant to genomic control.

The reaction opposite to that effected by HATs is cat-alyzed by histone deacetylases (HDACs). The history oftheir discovery begins, once again, in the mid-1980s, whena screen in budding yeast performed in the laboratory ofR. Gaber identified a number of loci that reverse a potas-sium transport deficiency (=RPD) by virtue of the fact thatmutations in those loci lead to the upregulation of potas-sium transporters. One such locus, RPD3, was subjectedto additional genetic analysis and found to be involved inthe control of transcription of a number of yeast genes.In 1996, the laboratory of S. Schreiber used an ingeniouspurification strategy to isolate a mammalian HDAC: theseresearchers reasoned that a competitive inhibitor could beused as bait for an HDAC (such a compound binds to thecatalytic site of an enzyme but prevents catalysis due toits structural difference from the enzyme’s bona fide sub-strate). Thus, the peptide trapoxin was used to preparean affinity matrix through which crude mammalian cellextract was passed; two polypeptides were found to asso-ciate with the matrix—and peptide microsequence analy-sis of one of them revealed its close sequence similarity tobudding yeast Rpd3p. Because activity assays indicatedthis newly purified protein to have HDAC activity, thesedata were accepted as the first evidence that an HDACin mammals may be involved in transcriptional control.Subsequent analysis of mammalian RPD3 (also known asHDAC1) confirmed that notion.

A dedicated effort to clone additional HDACs from eu-karyotic genomes revealed at least eight distinct genes—itis very likely that more will be identified once the emergingsequences of metazoan genomes, nematode, insect, rho-dent, and primate, are analyzed in sufficient detail. Severalgeneral observations can be made at this time, however.From a functional standpoint, many of the HDACs havebeen functionally connected to transcriptional repressionpathways; two examples are informative.

The first one involves one of the strongest mechanismsfor effecting transcriptional repression currently known:DNA methylation. Genomes of higher vertebrates con-tain unusually low amounts of the dinucleotide CpG (the-oretical predictions suggest each dinucleotide should ac-count for 1/42 = 0.0625, i.e., ca. 6% of the genome; CpG

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is significantly less abundant). In fact, the bulk of CpGdinucleotides occur within short (ca. 100–500 base pairs)stretches of the genome designated “CpG” islands—theseoccur most commonly in the promoters of genes. In ad-dition to this peculiar distribution, the cytosine withinCpG dinucleotides is frequently covalently modified witha methyl group on position 5 (to yield 5-methylcytosine,5-mC). There is very strong evidence that DNA methy-lation is a mechanism for targeting the transcriptionalrepression machinery to particular DNA sequences: forexample, in our genomes, the bulk of genomic parasites(i.e., transposons and retroviruses) are kept transcription-ally quiescent by methylation. Experiments indicate thatthe demethylation of repetitive DNA can have severe con-sequences for genomic stability: for example, retroele-ments begin uncontrolled proliferation in the genomes ofinterspecific hybrids in wallabies, and mutations in theenzyme that effects DNA methylation (DNA methyltrans-ferase) in humans leads to marked chromosome instabilitydue to pericentric heterochromatin expansion (this geneticdisorder—ICF syndrome—is characterized by skeletal ab-normalities and mental retardation). In addition, a verycommon feature of neoplasia in humans is the aberrantsilencing of genes required for cell cycle arrest (such asgenes for cyclin-dependent kinase inhibitors); this is ef-fected by hypermethylating the CpG islands in the pro-moters of these genes.

A functional connection between DNA methylation andthe targeting of histone deacetylase emerged from stud-ies on transcriptional regulators discovered in A. Bird’slab that bind methylated DNA selectively (i.e., these pro-teins bind 5mCpG but do not bind CpG) and appear to betranscriptional repressors. Subsequent biochemical anal-ysis in A. Bird and A. Wolffe’s lab revealed that onesuch protein—MeCP2—associates with histone deacety-lase and that an HDAC inhibitor prevents transcriptionalrepression driven by MeCP2. These observations illumi-nated the fact that hypermethylated DNA sequences re-side in deacetylated chromatin in vivo: thus, a simplemodel emerges according to which methylated DNA re-cruits specific proteins that recognize it selectively, andtarget HDAC to remodel chromatin adjacent to their bind-ing site into a repressive, deacetylated conformation. Theprofound relevance of this pathway to genomic control invivo can be seen from clinical and genetic evidence onhuman patients with mutations in the gene for MeCP2;these individuals develop Rett syndrome—a progressive,debilitating neurological disorder.

The second example regarding HDAC targeting in tran-scriptional repression involves certain members of the nu-clear hormone receptor (NHR) superfamily. Some of theseproteins—GR, for example—are constitutively cytoplas-mic and translocate to their site of action, the nucleus, only

in the presence of ligand. In contrast, other NHRs, includ-ing the retinoic acid receptor and the thyroid hormonereceptor, reside in the nucleus irrespective of hormone.Their residence in the nucleus is not a passive one—bothTR and RAR remain bound to target gene promoters inthe absence of hormone and act as potent transcriptionalrepressors. The exact mechanism whereby these proteinseffect repression remained elusive until work in 1995 fromD. Moore, R. Evans, and M. Rosenfeld identified twolarge (ca. 270 kDa) related polypeptides called N-CoR andSMRT. Subsequent work from these labs, and also those ofM. Lazar and A. Wolffe revealed unliganded TR and RARcan associate with HDAC via these corepressors, and thatHDAC inhibitors will impair the repressive action by theseregulators (the actual enzyme recruited by these proteinsis HDAC3). Whether the targeting of HDAC leads to chro-matin deacetylation of any vertebrate genes is not known.In budding yeast, work from the laboratories of M. Grun-stein and K. Struhl showed (via the application of ChIP)that the transcriptional repressor Ume6p acts to silencetarget genes via the recruitment of the HDAC Rpd3p, andthat such recruitment leads to histone tail deacetylationover target gene promoters.

The action of HDACs transcriptional repression path-ways is not an academic issue, since HDAC inhibitors havelong been known to be very potent cytostatic and differ-entiating agents. Thus, for example, a number of cell linesof cancerous origin can be driven to cease proliferationvia the application of such HDAC inhibitors as tricho-statin A or sodium butyrate. In addition, oncoproteins thatare produced as a result of chromosomal translocationsin leukemias are known to depend on HDAC targetingfor action; compounds that force a release of HDAC fromthese chimeric proteins are successfully used in clinicalpractice to treat certain forms of leukemia.

An interesting property of HDACs is their tendency tooccur in large, multisubunit complexes. For example, asoriginally discovered in the laboratory of A. Wolffe, andsubsequently by the laboratories of S. Schreiber and D.Reinberg, the predominant biochemical form of HDAC1in our cells is the Mi-2/NRD complex; its most distin-guishing feature is that it contains a histone deacetylase,a SWI/SNF family ATPase, and a protein that can selec-tively bind to methylated DNA. An understanding of thefunctional relevance of these associations awaits futherexperimentation.


Scientists that devoted their careers to the study of chro-matin are understandably pleased with the progress of the

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past decade: not only is our structural understanding ofchromatin (at least on its elementary level) more pro-found than it has ever been but also a large number ofprotein complexes has been discovered that are intimatelyinvolved in transcriptional control and that can remodeland modify chromatin structure. These developments aresomewhat of a mixed blessing, however, because the abun-dance of these protein factors, and our less-than-completeunderstanding of their in vivo function complicates at-tempts to depict gene activation and repression as a sim-ple, linear sequence of events (e.g., “protein binds to DNA,protein recruits RNA polymerase, RNA polymerase syn-thesizes mRNA”).

Two major questions in the field currently lack a com-prehensive answer: (i) what are the in vivo structural con-sequences of chromatin remodeling and modification interms of the behavior of the transcriptional machinery?;(ii) what is the relative contribution that chromatin mod-ification/disruption, and non-chromatin-based regulatorypathways make to gene activation and repression in vivo?While comprehensive answers are currently lacking, an at-tempt at a synthesis of the current data can be made, how-ever, based on experiments on the budding yeast HO en-donuclease gene (K. Nasmyth), MMTV LTR (G. Hager),mouse serum albumin gene enhancer (K. Zaret), the Xeno-pus TRβA gene (A. Wolffe), and the Gal4-VP16 activator(A. Belmont). This hypothetical scenario for gene acti-vation is presented in the form of a numbered list, butwe emphasize that the sequence of some steps may bechanged on specific promoters, and that some steps maybe omitted altogether.

1. A transcriptional regulator accesses its binding sitewithin a target promoter assembled into a maturenucleosomal array.

2. The chromatin-bound regulator targets anATP-dependent chromatin remodeling engine such asSWI/SNF; this targeting leads to the localizedremodeling of the histone DNA contacts and thegeneration of a DNAse I hypersensitive site.

3. This remodeling allows access to the promoter ofother nonhistone factors; in concert with the “pioneerfactor,” these target HAT-containing complexes; theiraction may promote the large-scale unfolding ofchromatin, as well as exert some local effect.

4. Some of the transcriptional regulators bound to thepromoter use other adapter complexes to promote theassembly of the preinitiation complex and thetargeting of RNA polymerase.

5. A number of complex events lead stalled RNApolymerase to begin productive transcriptionalelongation (which may be facilitated byhyperacetylation).

It is very clear that transcriptional control harbors path-ways and mechanisms that “are not dreamt of” in ourcurrent paradigms. The integration of chromatin infras-tructure into gene expression regulatory pathways, how-ever, is fairly certain to remain at the core of our notionsof genome control.





Bird, A. P., and Wolffe, A. P. (1999). “Methylation-induced repression—belts, braces, and chromatin,” Cell 99, 451–454.

Elgin, S. C. R. (ed.). (1995). “Chromatin Structure and Gene Expression,”IRL Press, Oxford.

Grunstein, M. (1997). “Histone acetylation in chromatin structure andtranscription,” Nature 389, 349–352.

Kingston, R. E., and Narlikar, G. J. (1999). “ATP-dependent remodelingand acetylation as regulators of chromatin fluidity,” Genes Dev. 13,2339–2352.

Lemon, B., and Tjian, R. (2000). “Orchestrated response: A symphonyof transcription factors for gene control,” Genes Dev. 14, 2551–2569.

Luger, K., Mader, A. W., Richmond, R. K., Sargent, D. F., and Richmond,T. J. (1997). “Crystal structure of the nucleosome core particle at 2.8A resolution,” Nature 389, 251–260.

Ng, H. H., and Bird, A. (2000). “Histone deacetylases: Silencers forhire,” Trends Biochem Sci. 25, 121–126.

Robertson, K. R., and Wolffe, A. P. (2000). “DNA methylation in healthand disease,” Nature Rev. Genet. 1, 11–19.

Sterner, D. E., and Berger, S. L. (2000). “Acetylation of histones andtranscription-related factors,” Microbiol. Mol. Biol. Rev. 64, 435–459.

Struhl, K. (1998). “Histone acetylation and transcriptional regulatorymechanisms,” Genes Dev. 12, 599–606.

Sudarsanam, P., and Winston, F. (2000). “The Swi/Snf family:Nucleosome-remodeling complexes and transcriptional control,”Trends Genet. 16, 345–351.

Urnov, F. D., and Wolffe, A. P. (2001). “A necessary food; Nuclear hor-mone receptors and their chromatin templates,” Mol. Endo. 15, 1–16.

Wolffe, A. P. (1998). “Chromatin Structure and Function,” AcademicPress, San Diego, CA.

Wolffe, A. P., and Hayes, J. J. (1999). “Chromatin disruption and modi-fication,” Nucleic Acids Res. 27, 711–720.

Wolffe, A. P., and Matzke, M. A. (1999). “Epigenetics: Regulationthrough repression,” Science 286, 481–486.

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DNA Testing in Forensic ScienceMoses S. SchanfieldMonroe County Public Safety Laboratory

I. What Is DNA?II. Extraction of DNAIII. Quantitation of DNAIV. Current Forensic DNA TestingV. STRs Used Forensically

VI. PCR Setup and DetectionVII. Usefulness in Detecting MixturesVIII. Individualization


Alleles Alternate forms of an inherited trait. Such as the“A,” “B,” or “O” alleles of the ABO blood group. Sim-ple polymorphisms will have as few as two alleles,while hypervariable polymorphisms will have five ormore alleles, such that the majority of the populationis heterozygous.

Amplified fragment length polymorphisms (AFLPsor AmpFLPs) Polymorphism in length of DNA seg-ments detected using the PCR process to amplify aspecific segment of DNA, followed by separation us-ing electrophoresis. These are normally Variable Num-ber Tandem Repeat (VNTR) regions that can havedifferent size repeats. AFLPs are classified by thesize of the repeat, repeats consisting of 10 or morebases are referred to as large tandem repeat (LTR)AFLP, while those containing seven or fewer base

“Mentioning of commercial products is not an endorsement by TheMonroe County Public Safety Laboratory.”

pairs per repeat are referred to as short tandem repeats(STRs).

Gene A sequence of DNA which codes for the productionof a specific protein or part of a specific protein, such asa glactosyl transferase (the B allele of the ABO bloodgroup).

Genotype The genetic type of a person determined ei-ther by the fact that they have two different alleles at alocus, or that the individuals parents have been testedto determine what alleles the person has geneticallyinherited.

Heterozygotes An individual with two different alleles ata given locus.

Homozygotes An individual with two alleles that are thesame.

Locus The place (location) where a specific gene residesin the human genome. The names of genetic regionshave been standardardized by the International Systemof Gene Nomenclature (Shows et al., 1987). Inheritedtraits that code for genes usually have a name that re-flects its biological function, such as THO1 for the


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first tyrosine hydroxylase locus. Some identified in-herited traits do not code for proteins, but are regionsthat show variation (polymorphism) in length or se-quence of DNA. These areas are referred to as “DNAloci,” such as D1S7, which represents and DNA inher-ited trait (D), located on chromosome 1, “S” indicatesit is a single copy region and “7” indicates it was theseventh polymorphism found on that chromosome.

Phenotype The observed results of a genetic test. If a per-son has two different alleles (e.g., is heterozygous), thephenotype and the genotype are the same. However, ifa person only has one allele detected, without testingthe parents we cannot be certain that the person hastwo alleles the same (i.e., homozygous) since the per-son could have two alleles the same, or one detectedallele and one undetected allele, for what ever reason.In the case of RFLP loci this is usually referred to asa single band pattern. For AFLP- and sequence-basedPCR-based systems this is referred to as a homozygousphenotype. In general since we do not know the genetictype of homozygous individuals it is better to refer tohomozygous and heterozygous phenotypes, unless thegenetic type of the individual is determined by pedigreeanalysis.

Polymorphism Genetically inherited variation with twoor more forms, the least common of which occurs at afrequency of greater than 1%.

Restriction fragment length polymorphism (RFLP)Polymorphism in length of DNA segments detected us-ing a restriction enzyme, followed by separation usingelectrophoresis.

Variable number tandem repeat (VNTR) regions of in-herited variation that consist of alleles which containdifferent numbers of repeating segments. It is easiest tothink of them as freight trains with different numbersof box cars. Different loci will have different numbersof repeats. Though RFLP loci are also VNTR loci thenumber of repeats is generally not known because ofthe detection technology.

FORENSIC SCIENCE is applied science. That is to saythat the scientific methodologies used in forensic sciencewere developed by biologists, chemists, and geneticistsand then taken over by forensic scientists to help themsolve problems. The first inherited trait to be used in foren-sic testing was the ABO blood group on red blood cellsand secreted blood group substance found in saliva andother body fluids of individuals called “Secretors.” Re-search in the 1950s, 1960s, and 1970s identified proteinsthat had genetic variation or were “polymorphic.” Someof these were enzymes such as acid phosphatase [ACP,referred to as erythrocyte acid phosphatase (EAP) by

forensic scientists], esterase D (ESD), phosphoglucomu-tase (PGM1) and some transport and functional proteinssuch as group specific component [GC, now known asVitamin D binding globulin [VDBG]], haptoglobin (HP),the immunoglobulin allotypes (GM and KM), and trans-ferrin (TF) were used forensically. These markers wereused in the late 1970s and early 1980s by forensic sciencein the United States and abroad to individualize bloodstains. Although, some of these markers did not last longin bloodstains they made it possible to often individual-ize bloodstains with greater than a 99% certainty. Semenstains, saliva, and urine had relatively few markers thatcould be detected, making it difficult to provide informa-tion in cases with these types of evidence.

In the mid-1970s two independent areas of researchwould change the future of forensic science. Research onbacterial enzymes that cut DNA at specific places led tothe development of restriction fragment length polymor-phisms (RFLPs). Initially these were only useful for thediagnosis of genetic diseases such as Sickle Cell Anemiacaused by a mutation in the hemoglobin gene. In 1980with the identification of the first hypervariable DNA poly-morphism (D14S1), detected by restriction length poly-morphism technology (RFLP), the door was opened tothe possibility that DNA technology could be applied toforensic evidence. In the next several years, the search fornew markers lead to the identification of many forensi-cally useful markers, detected by RFLP, some of which arestill used routinely in forensic DNA testing. In the mean-time, hypervariable minisatellite regions which identifiedmany genetic regions at one time (multilocus probes) werefound. The term “DNA fingerprinting” was used to de-scribe these bar code-like patterns. Although these regionsproved to be highly informative for parentage testing, theydid not have the sensitivity needed for forensic testing.Though multilocus probes were used for paternity testingand some forensic applications, they are rarely used at thepresent time. Using a battery of five to seven RFLP locimade it possible to individualize samples into the 100s ofmillions and billions. This means that the chance of anytwo unrelated individuals matching was very unlikely.

At the same time the revolution in RFLP was beginningin the mid 1970s, an early version of copying DNA usingrepair enzymes called polymerases was being explored.It would not be until the early 1980s when the modernPolymerase Chain Reaction (PCR) tests was developed.The role of the polymerase enzymes in copying DNA hadbeen known since the 1970s. The use of high temperatureTaq polymerase allowed for the automation of thermalcycling and the introduction of modern PCR. Tests weredeveloped to identify human leukocyte antigens (HLA) forthe transplantation community. The first marker that theydeveloped was to the HLA region called DQα (now called

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DQA1). This marker looked at a genetic marker that hadoriginally been tested at the protein level at the DNA level.This type of marker looked at DNA-sequence-based dif-ferences. This became the first PCR based tested to be usedforensically. Other sequence-based tests were developedbut they did not provide the same level of identificationproduced by RFLP based testing. Using the available se-quence based tests only allowed individualization in the100s to 100,000s.

The FBI established a standardized system for publiclyfunded crime laboratories in the United States. Similarwork was going on in Canada, the United Kingdom, andEurope. The introduction of DNA restriction fragmentlength polymorphism (RFLP) technology revolutionizedthe field of forensic identity testing. This is especiallytrue for the area of sexual assault evidence that histori-cally has been an area of limited information. With DNA-based testing sperm DNA could be separated from thevictims type allowing for the first-time regular direct test-ing of the sperm donor. Though RFLP technology hasbeen a tremendous aid, it has several problems. It is ex-pensive to implement, labor intensive, expensive to test,and is limited by both quantity and quality of DNA ob-tained. Further, because the process involved measuringthe movement of bands and not directly the DNA productthere were many statistical problems with representing thedata. The technical feasibility of amplifying specific seg-ments of DNA using the polymerase chain reaction (PCR)had the potential to overcome the shortcomings of RFLPtechnology.

PCR-based technology is much less expensive to imple-ment, since it does not require a laboratory capable of han-dling radioactive isotopes. It has higher through put sinceeach worker can do more cases in the same amount of time.PCR by its nature works with smaller amounts of DNAand with DNA that has been environmentally abused. Fi-nally, since the DNA product can be identified as differentalternative forms the statistical manipulation of data wassimilar to that for other genetic polymorphisms.

In 1989 at the same time that the RFLP-based testingwas being converted to PCR-based testing creating the firstamplified fragment length polymorphism or AFLP, newpolymorphisms were being found directly using PCR. Theconverted RFLP loci which consisted of different num-bers of repeated segments, much like freight trains withdifferent numbers of box cars were referred to as “variablenumber of tandem repeat” or VNTR regions or loci. Theserepeats consisted of 15 to 70 base pairs and were referredto at ”large tandem repeat loci or LTRs. The new regionsbeing found had much smaller repeats consisting of two,three, four, or five bases in a repeat unit. These new mark-ers were called “short tandem repeats” or STRs for short.The four base pair or tetra nucleotide repeats became the

genetic markers used to map the human genome. Withthis large number of markers available it became possibleto pick sets of these markers to make highly discrimina-tory multiplexes (multiple regions amplified at the sametime). The use of fluorescent detection of DNA fragmentson automated DNA sequencers from the human genomeproject made it possible to create fluorescent multiplexeswith automated detection. These methodologies are nowthe methods of choice for use in the forensic testing ofDNA samples.


DNA stands for deoxyribonucleic acid. It is the biologicalblueprint of life. DNA is made up of a double-strandedstructure consisting of sugar (deoxyribose) and phosphateback bone, cross linked with two types of nucleic acids re-ferred to as purines (adenine and guanine) and pyrimidines(thymine and cytosine) (Fig. 1). The cross linking nucleicacids always pair a purine with a pyrimidine, such thatadenine always pairs with thymine and guanine alwayspairs with cytosine.

DNA can be found in several areas of a cell. The ma-jority of DNA is located in the nucleus of cells (Fig. 2)organized in the form of chromosomes (22 pairs of auto-somes and a set of sex chromosomes (X and Y)). Eachnucleated cell normally has 46 chromosomes that repre-sent the contribution from both parents. In the formationof gametes (eggs and sperm) one chromosome of each pairis randomly separated and placed in the gamete. The sep-aration of chromosomes is referred to as segregation. Thetransmission of half of our chromosomes to our children inthe form of gametes is the basis of Mendelian inheritance.This DNA is referred to as nuclear or genomic DNA. Withthe exception of identical twins, no two people share thesame genomic DNA sequence.

Another source of DNA is found in the mitochondriain the cytoplasm of cells (Fig. 2). Unlike nuclear DNA,which only has two copies of each genetic region, mito-chondrial DNA is involved in energy production withinthe cell and can have between 100 and 10,000 copies percell. Structurally, instead of a linear arrangement of DNAwithin chromosomes, mitochondrial DNA has a circularstructure. Mitochondrial DNA is inherited from the motherbecause it is found in the cytoplasm which comes from theegg (ova).

A. Where Is DNA Found?

Nuclear or genomic DNA is found in all nucleated cellsas well as in the reproductive cells (eggs and sperm). Theamount of DNA we can expect to find in different cellsand types of evidence are found in Table I. DNA has been

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FIGURE 1 Molecular structure of DNA. From top to bottom: Adenine–Thymine, Guanine–Cytosine, Adenine–Thymine and Guanine–Cytosine. (From Schanfield, M. S. (2000). Deoxyribonucleic Acid/Basic Principles. In“Encyclopedia of Forensic Sciences” (Siegel, J. A., Saukko, P. J., and Knupfer, G. C., eds.), Academic Press, London,p. 481.)

successfully obtained from blood and bloodstains, vagi-nal and anal swabs, oral swabs, well-worn clothing, bone,teeth, most organs, and to some extent urine. It is less likelyto obtain DNA from some types of evidence than others.Blood or semen stains on soil and leather are historicallynot good sources of evidenciary DNA. Saliva, per se, hasfew nucleated cells, but, beer and wine bottles, drinkingglasses, beer cans, soda cans, cigarettes, stamps and enve-lope flaps have all been found to provide varying amountsof DNA.

B. How Much DNA Is Neededfor Forensic Testing?

The amount of DNA needed to perform testing dependson the technology used. RFLP technology usually needs atleast 50 ng of intact high-molecular-weight DNA. In con-trast PCR-based testing can use as little as 250 pg. Most

PCR based tests are set up to use between 1 and 10 ng ofgenomic DNA.

C. Destruction of DNA

Biological materials are going to be affected by their en-vironment. Enzymes lose activity over time and type ofstorage conditions. DNA has been found to be relativelyrobust when it is in the form of dry stains. Initial envi-ronmental studies indicated some of the limitations ofDNA based on the material it is deposited upon and theenvironmental conditions. Environmental insult to DNAdoes not change the results of testing, you will either ob-tain results, or if the DNA has been too badly affectedby the environment (i.e., the DNA is degraded) you donot get RFLP results. One report on the success rate ofobtaining RFLP results and noted that depending on thesubstrate or condition of the stain, results were obtained

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FIGURE 2 A generalized eukaryotic cell showing the organization and distribution of organelles as they would appearin transmission electron microscope. The type, number, and distribution of organelles are related to cell function. (FromSchanfield, M. S. (2000). Deoxyribonucleic Acid/Basic Principles. In “Encyclopedia of Forensic Sciences” (Siegel, J. A.,Saukko, P. J., and Knupfer, G. C., eds.), Academic Press, London, p. 481.)

between 0 (carpet stains or putrefied samples) and 61.5%(scrapped dried stains) of the time with and average of52% for the 100 items of evidence tested.. Thus, the ma-terial DNA is deposited on and the degree of further in-sult can markedly affect the ability to obtain RFLP DNAresults.

All of the published studies on environmental insultwere done on prepared dried stains. Since biological flu-ids are liquid the effects of ultraviolet radiation on liquidDNA have been evaluated. The results of exposing 100-µlsamples of a standard DNA solution to fluorescent lightin the laboratory, a UV germicidal light (254 nm), mid-day sunlight in January, and early sunset light in January in

TABLE I DNA Content of Various Tissues

1 sperm 3 pg

1 cell 6 pg

1 shed hair 1 nga

1 plucked hair 300 ngb

1 drop of blood 1,500 ng

a The success rate for PCR on shed hairs is 30 to 50%,so this average is optimistic.

b There is a great deal of variation among hair roots.Fine blond hair will tend to have much less DNA, whilecourse dark hair with large roots more.

15-min increments, up to 1 hr are presented in Fig. 3. Thereis a linear decrease in high-molecular-weight DNA withthe UV germicidal light, such that after an hour about 96%of the high-molecular-weight DNA has been lost. Even inthe weak midday light in January, over 60% of the high-molecular-weight DNA was lost. In contrast, the fluores-cent lighting in the laboratory and the after sunset light had

FIGURE 3 Plot of DNA concentration (frorescence) over timeafter exposure to different light sources. (From Schanfield, M. S.(2000). Deoxyribonucleic Acid/Basic Principles. In “Encyclopediaof Forensic Sciences” (Siegel, J. A., Saukko, P. J., and Knupfer,G. C., eds.), Academic Press, London, p. 482.)

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no effect on the amount of high-molecular-weight DNA.This was not a rigorous experiment, but the effects aredramatic enough to demonstrate the effect of ultra violetlight exposure to DNA before stains dry.


As stated previously DNA exists inside of cells. Becausemost evidence is in the form of dry stains, the DNA mustbe removed from the stain before it can be tested. Theprocess of removing DNA from the cells on the evidenceand dissolving it is referred to as extraction. There areseveral procedures available for removing DNA from ev-idence so that it can be used. They are referred to as ei-ther “organic” extraction or “nonorganic” extraction basedon the nature of the chemicals used. Further, there aretwo special types of extraction. The first, called differ-ential extraction, was developed for sexual assault ev-idence to separate the cells that come from the victim(epithelial cells from the vagina, rectum, or mouth) fromthose of the perpetrator (male sperm cells). The secondmethod is a specialized “nonorganic” extraction usingChelex beads. Chelex beads can only be used when PCR-based DNA testing is going to be used. The basic DNAextraction procedures, whether organic or nonorganic,can be adapted for special circumstances such as hair ortissue.

A. Chloroform–Phenol Extraction

This is the oldest procedure available for extracting DNAfrom blood and it has been extended to include hair, tissue,and semen stains. The basic procedure consists of open-ing up cells with a buffer and an enzyme, usually Pro-tease K, and then denaturing and separating the proteins

TABLE II Comparison of Organic and Nonorganic Extraction of DNA fromBlood and Semen Stainsa

Quantitation Blood Blood Semen Semenmethod organic nonorganic organic nonorganic

Yield gel

Mean 185 ng 258 ng 175 ng 207 ng

N 21 8 22 8

p .054 .122

Slot blot

Mean 515 ng 908 ng 627 ng 1175 ng

N 22 8 27 8

p .022 .008

a Data taken from Tables 1 and 2 of Laber et al. (1992). Differences in meanstested by Kruskall–Wallace nonparametric analysis of variance, H statistic with 1 df,uncorrected p values presented.

from the DNA. The latter part is done using a mixture ofchloroform (24:1 chloroform:isoamyl alcohol) and phe-nol (buffered). The phenol–chloroform mixture denaturesproteins liberated by the first stage. The major disadvan-tage of this procedure is the fact that phenol–chloroformis a hazardous waste and could theoretically pose a riskto pregnant employees. A modern protocol for phenol–chloroform extraction of various types of evidence can befound in the literature.

B. Nonorganic Extraction

In nonorganic extraction the hazardous phenol–chloro-form protein denaturation step was replaced by a saltingout of proteins. This allowed for the same chemistry tobe used for the initial phase of DNA extraction, and re-placement of the hazardous elements of the procedurewith a nonhazardous alternative. The salting-out proce-dure has several advantages over the phenol–chloroformextraction. The first is that instead of having two liquidphases (organic and nonorganic) that can occasionally trapthe DNA in the wrong phase (organic) phase, by precip-itating the proteins (e.g., the proteins become insolubleand become a solid), there are liquid and solid phaseswith the DNA only in the liquid phase (nonorganic).The second advantage is that the hazardous phenol–chloroform is replaced with a harmless salt solution. Com-parison of the organic and nonorganic procedures forblood, and semen indicate that the nonorganic extrac-tion is on the average as good or better than organicextraction, whether quantitated by yield gel or slot blot(Table II).

Either method of DNA extraction described earlier canbe used for both RFLP- or PCR-based DNA testing. Or-ganic DNA extraction is widely used in laboratories doingcriminal casework while nonorganic DNA extraction is

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widely used in laboratories performing paternity testing,research, and diagnostics. On a worldwide basis nonor-ganic DNA extraction is the more prevalent. With the shiftto PCR-based testing this choice in extraction is increas-ingly common.

C. Chelex Extraction

In 1991, a method of DNA extraction was described thatwas specifically aimed at the extraction of small amountsof dilute DNA for PCR-based testing using Chelex beads.The method is simple, relatively fast, and biohazard free. Itis widely used by forensic laboratories doing PCR-basedtyping which has increased the number of laboratories us-ing nonorganic, biohazard-free DNA extraction. The onlylimitations of Chelex extraction is that it produces a dilutesolution of DNA that may need to be concentrated be-fore it can be used with some of the newer high-resolutionPCR-based typing systems and it cannot be used for RFLPtesting.


A. Yield-Gel Quantitation

Whether RFLP- or PCR-based testing is performed it isnecessary to know how much DNA is present. One of theearliest methods of quantitating small amounts of DNAis the use of a yield gel. A small gel is made using asalt solution to carry electrical current and a supportingmedium made of agarose (a complex carbohydrate madefrom seaweed). Much like gelatin, the agarose dissolvesin water that is heated to near boiling and the liquid iscooled slightly and poured into a casting tray. A plasticcomb or well former with rectangular teeth is placed inthe liquid agarose. Once the agarose gels, the comb isremoved leaving behind rectangular wells in the agarosegel. The DNA to be tested is mixed with loading buffer,and placed in the wells. Loading buffer is a mixture of alarge amount of sugar and dye. The high concentration ofsugars makes the mixture heavier than the salt solutionso that the DNA sinks to the bottom of the well. Thedye allows the migration of the DNA to be monitored.The agarose was melted in water containing salt. Whenelectrical current is applied to the gel, electricity flowsthrough the gel because of the salt and moves (migrates)from the negative electrode (cathode) toward the positiveelectrode (anode). Since all DNA has a negative charge,and was placed in the wells at the cathodal end of the gel,the negatively charged DNA will migrate out of the wellstoward the positive end of the gel. If the DNA is brokeninto pieces that are different sizes, the smaller pieces willmove through the gel faster than the larger pieces and will

be separated based on size. This process of separatingDNA using an electric current is called electrophoresis,which simply means separation (phoresis) by means ofelectricity (electro).

Since DNA is colorless it is not possible to see the DNAafter it has been separated without the use of special dyesthat bind to it. One of the earliest dyes used was ethidiumbromide, which fluoresces pink when bound to double-stranded DNA and exposed to ultraviolet light. Figure 4 isan ethidium bromide stained yield gel. To quantitate theamount of DNA in the DNA extracts, a set of DNA quan-titation standards are placed on the gel. By visual compar-ison of the unknown DNA to the known DNA the amountof DNA can be approximated. This test provides informa-tion about the relative amount of DNA and whether it isdegraded (i.e., the DNA is broken down so that different-size pieces of DNA are present). It does not indicate if theDNA is human, however, since all DNA will fluoresce.Thus the DNA present may be bacterial as well as hu-man DNA. For RFLP testing the total amount of DNA inthe sample is the important determinant of how the sam-ples migrate in the gel. Therefore yield-gel electrophoreticquantitation of DNA is an appropriate method. Yield-gelquantitation of DNA for RFLP testing was considered tobe such an integral part of quality assurance that it wasincluded in the National Institute of Standards, StandardReference Material 2390, “DNA Profiling Standard.”

As with the extraction of DNA using the organicmethod, ethidium bromide is potentially hazardous be-cause the dye is associated with an increased cancer risk.Though ethidium bromide is still widely used for the iden-tification of DNA it is currently being replaced by a newdye called Sybrr® green which is much less carcinogenicand can detect smaller amounts of DNA than ethidiumbromide.

B. Slot-Blot Quantitation

In contrast to RFLP, for PCR-based testing, the amountof human DNA and not the total amount of DNA is animportant determinant in how likely it will be to obtainresults. A slot blot does not rely on electrophoresis to sep-arate the DNA but rather on the ability of denatured (sep-arated DNA strands) DNA to bind to homologous com-plementary sequences. The ability to quantitate humanDNA requires sequences of DNA that are common in thehuman genome so that a single DNA sequence can recog-nize them and bind to them. The repeated DNA sequencecalled D17Z1 is the basis for all human DNA slot-blotquantitation systems. There are several of these proce-dures commercially available. In one of the most widelyused tests, the quantitation requires that denatured DNAis applied to a membrane using a slotted plastic appa-ratus. The denatured DNA binds to the membrane. The

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FIGURE 4 Ethidium bromide stained yield gel. Bottom left samples are quantitative standards. Other samples repre-sent various samples of DNA. Upper right sample is degraded DNA. (From Schanfield, M. S. (2000). DeoxyribonucleicAcid/Basic Principles. In “Encyclopedia of Forensic Sciences” (Siegel, J. A., Saukko, P. J., and Knupfer, G. C., eds.),Academic Press, London, p. 484.)

membrane is exposed to a solution of denatured DNAfragments that recognizes a repeating sequence of humanor primate DNA. Pieces of DNA that recognize a specificregion of DNA are referred to as a “probe.” The probewill bind to complementary DNA fragments stuck on themembrane. The probe has an indicator attached to it sothat the binding of the DNA to the probe can be detected.The unbound probe is washed off and the probe is detectedusing either chemicals that change color (colorimetric de-tection) or chemicals that give off light (chemiluminscentdetection). To be able to quantitate the amount of humanDNA present, standards with different amounts of humanDNA are also placed on the membrane so that it is possibleto determine the approximate amount of DNA bound to themembrane by visual comparison to the known standards.More precise quantitation can be obtained by scanningthe membrane with a scanning densitometer and deter-mining the amount of color associated with each band.Most forensic laboratories use visual comparison.


At this point in time, at the beginning of the 21st century,forensic DNA testing has moved away from RFLP test-

ing and is replacing sequence-based PCR strip technologywith fluorescent STR-based testing. Thus it is important tounderstand the nature of PCR-based testing and the powerit provides to forensic scientists.

A. Definition and Description of PCR

The Polymerase Chain Reaction (PCR) is based on bio-chemical processes within cells to repair damaged DNAand to make copies of the DNA as the cells replicate. Inthe repair mode, if a single strand of DNA is damaged,the damaged area is removed so that there is a single-stranded section of DNA with double-stranded sectionsat either end. The polymerase enzyme fills in the missingcomplementary DNA. In the copy mode an entire strandis copied during DNA replication. Figure 5 illustratesa polymerase enzyme copying a portion of a strand ofDNA.

In a cell a specific gene is copied or translated from DNAto RNA because the polymerase has specific start and stopsignals coded into the DNA. To copy a sequence of DNAin vitro, artificial start and stop signals are needed. Thesesignals can only be made once the sequence of the re-gion to be amplified is known. Once a sequence is known,the area to be copied or amplified can be defined by a

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FIGURE 5 DNA polymerase copying one strand of a portion of double-stranded DNA. (From Schanfield, M. S. (2000).Deoxyribonucleic Acid/Polymerase Chain Reaction. In “Encyclopedia of Forensic Sciences” (Siegel, J. A., Saukko,P. J., and Knupfer, G. C., eds.), Academic Press, London, p. 516.)

unique sequence of DNA. For a primer to recognize aunique sequence in the genome it must be long enoughfor no other sequence to match it by chance. This can usu-ally be achieved with a sequence of 20 to 25 nucleotides.These manufactured pieces of DNA are called “primers”and they are complementary to the start and stop areasdefined earlier. The “forward primer” is complementaryto the beginning sequence on one strand of DNA, usuallycalled the positive strand. The “reverse primer” is comple-mentary to the stop sequence on the opposite or negativestrand of DNA.

B. Multiplexing PCR Reactions

One of the advantages of PCR is that more than one regioncan be amplified at a time. Although it is necessary toselect carefully primers that cannot bind to each other, theonly limitation is how many pairs of primers can be placedtogether is the ability to detect the amplified product.

C. PCR Process

To perform a PCR reaction several ingredients are needed.They include PCR reaction buffer, which is basically a saltsolution at the right pH for the enzyme being used, thefour nucleotides (DNA building blocks), primers, a ther-mostable DNA polymerase (Taq, Pfu, Vent, Replinase,etc), and template DNA. These reactants are placed insmall plastic reaction tubes. The process consists ofheating a solution of DNA to greater than 90◦C. Double-stranded DNA comes apart or melts to form single-stranded DNA at this temperature. This is called the denat-uration step. The solution is then cooled down to between50 and 65◦C the primers will bind to their complemen-tary locations. This is called the annealing or probe hy-bridization step. Finally, the solution temperature is raisedto 72◦C at which point the polymerase makes a copy of

the target DNA defined by the primers. This is called theextension step. This completes one cycle of the PCR pro-cess. To make enough copies of the target DNA to detectthe process is repeated from 25 to 40 times. This is doneusing a device called a thermalcycler. The process is illus-trated in Fig. 6. If the process were perfect 30 cycles wouldcreate over a billion copies of the original target DNA.

The heating and cooling of the tubes are done in anelectromechanical device call a “thermalcycler,” which ingeneral, consists of an aluminum blocks with wells de-signed to fit the plastic PCR reaction tubes. The aluminumblock has heating and cooling elements controlled by a mi-croprocessor that can raise and lower the temperature ofthe block and the plastic PCR reaction tubes in the block.In the thermal cyclers that were first made, the plasticreaction tubes extended above the thermal block. This al-lowed cooling to take place above the reaction. The waterin the reaction mixture would evaporate and condense atthe top of the tube, changing the concentration of reac-tants and affecting the success of the amplification. Tolimit the evaporation, mineral oil was placed on top of thereaction mixture. New thermal cyclers have heated lids ontop of the block to prevent or minimize evaporation. Themicroprocessor can store many sets of instructions suchthat different programs can be kept in the microprocessorto amplify different sequences of DNA.

D. Detection of PCR Products

There are many methods for detecting PCR products.Since large amounts of product are produced there is noneed to use techniques such as radioactive detection, al-though it has been used in some clinical setting. In forensictesting, one of the advantages of PCR-based testing is thatit does not require the use of hazardous materials to detectit. There is normally enough product so that if the PCRproducts are run on a yield gel and stained with ethidiumbromide or Cyber green, there is normally enough DNA

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598 DNA Testing in Forensic Science

FIGURE 6 The PCR process. Courtesy of PE Cetus Instruments. (From Schanfield, M. S. (2000). DeoxyribonucleicAcid/Polymerase Chain Reaction. In “Encyclopedia of Forensic Sciences” (Siegel, J. A., Saukko, P. J., and Knupfer,G. C., eds.), Academic Press, London, p. 517.)

to be detected. This is a suggested method to verify if thePCR amplification was successful and that there is a PCRproduct to detect.


At this point in time with the large number of STR loci, ademand for a standardized panel in the United States, anda need for there to be at least some sharing of loci withforensic counterparts in Canada, England, and Europe,the Technical Working Group on DNA Analysis Methods(TWGDAM) implemented a multi-laboratory evaluationof those STR loci available in kits in the United States. Theloci chosen would be the PCR-based core of a national sexoffender file required under the 1994 DNA IdentificationAct. The national program is called Combined DNA In-dexing System or CODIS for short.

The TWGDAM/CODIS loci were announced at thePromega DNA Identification Symposium in the fall of1997 and at the American Academy of Forensic Sci-ence meeting in February 1998. The following loci werechosen to be part of what was originally called theCODIS 13 loci: CSF1PO, D3S1358, D5S818, D7S820,

D8S1179, D13S317, D16S539, D18S51, D21S11, FGA,THO1, TPOX, and VWA03. These loci overlapped withthe Forensic Science Services multiplexes and the Interpolmultiplexes. The 13 loci can be obtained in two amplifica-tions using Profiler Plus and Cofiler or in a single ampli-fication using a kit in development called Identifiler from

1 and Powerplex 2 from Promega (Table II), or in a singlereaction with Powerplex 16 by Promega.

A. Fluorescent Dyes

Before discussing the equipment used to detect fluorescentSTRs some understanding of fluorescent dyes are neces-sary. Fluorescent dyes or minerals when subjected to lightat one wave length, such as untraviolet light or black light,will give off colored light at a slightly different wavelengthor color. A characteristic of fluorescent dyes or materialsis that the compound is excited at one frequency of light,referred to as its absorption or excitation peak, and emitsor gives off light a different frequency, referred to as itsemission peak.

To label a PCR primer with a fluorescent dye, the dyeis attached to the 5′ end of the molecule. Since DNA is

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translated from 5 to 3′ it is at the very end of the primerand should not affect the amplification or binding of theprimer if made correctly. One of the oldest fluorescentdyes is fluoroscein. Many devices have been made thatwill detect fluoroscein, and it has been used extensively tolabel antibodies and other materials. Many of these dyeshave been used for a long time and are in the public do-main. Others have been developed for specific projects.The dyes used by PE Biosystems were originally propri-etary and part of a patented four color DNA sequencingsystem (Blue, Green, Yellow, Red). These dyes are nowbecoming more readily available.

B. Fluorescent Detection Equipment

The equipment used to detect the product of fluorescentlylabeled STR tests fall into two categories. The first are

FIGURE 7 (A) is the original CCD images of Powerplex 1.1 (Panels 1 and 2 ) and Powerplex 2.1 (Panels 3 and 4 ).(B) is the reverse image of the same images. (B) Panel 1 is the TMR loci from top to bottom CSF1PO, TPOX, THO1,and VWA03. (B) Panel 2 is the fluoroscein loci, from top to bottom D16S539, D7S820, D13S317, and D5S818. (B)Panel 3 is the TMR loci from top to bottom FGA, TPOX, D8S1179, and VWA03. (B) Panel 4 is the fluoroscein loci fromtop to bottom Penta E, D18S51, D21S11, THO1, and D3S1358. The same samples are loaded between the ladders.The sample format is the TWGDAM format when an internal size standard is not used. The overlapping loci for sampleconfirmation are THO1, TPOX and VWA03. (From Schanfield, M. S. (2000). Deoxyribonucleic Acid/Polymerase ChainReaction-Short Tandem Repeats. In “Encyclopedia of Forensic Sciences” (Siegel, J. A., Saukko, P. J., and Knupfer,G. C., eds.), Academic Press, London, p. 530.)

devices that scan a gel after the DNA products havebeen separated by a process called electrophoreses. Ex-amples of post electrophoresis scanners are the HitachiFMBIO® Fluorescent Scanner, the Molecular DynamicsFluorimagerTM and the Beckman Genomics SC scanner.The Hitachi and Molecular Dynamics use a laser as a lightwith filters to identify the proper frequency and a CCDcamera to capture the image. The Beckman Genomics SCuses a monochromatic Xenon light source and uses filtersto detect the appropriate light for the CCD camera. TheCCD camera scans back and forth over the gel as it is ex-posed to the light source and detects the various fluorescentcolors using filters that change. This type of equipmenthas flexibility because different formats of electrophore-sis gels can be used and scanned. The output is in the formof an electronic image with bands, that look much like a setof RFLP bands. Figure 7A is an example of actual images

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600 DNA Testing in Forensic Science

(light on dark) recorded by the CCD camera, and Fig. 7Bis the reverse image (dark on light) that is reminiscent ofan RFLP autoradiograph or lumigraph. It should be notedthat the scanning time is added onto the electrophoresistime, with increased time for each color read.

The second type of imaging system is a real time sys-tem, in which the DNA fragments, after the bands havebeen resolved, pass beneath a light source scanner that re-covers the spectrum of light from the different fluorophors.This is the ABI Prism® system from PE Biosystems. It in-cludes the older Model 373 DNA Sequencer and 377 DNASequencer, which use slab acrylamide electrophoresisto separate the DNA fragments, and the 310 and 3100Genetic Analyzers which use capillary electrophoresis toseparate the DNA fragments. Capillary electrophoresis isa technology in which a fine glass capillary is filled with aproprietary separation polymer. The sample is pulled intothe capillary by applying an electric current to it, then us-ing high-voltage electrophoresis (12,000 V), and the DNAfragments are separated over the length of the column andmove past a laser detector. The 377 can put approximately60 samples on a gel at one time, and with modifications, 96.In contrast, the 310 CE system does one sample at a time,with a separation time of approximately 20 min. However,as this is automated, a cassette can be filled with samplesfor testing and left to run unattended. The 3100 uses 10capillary tubes with higher throughput. The output of thesedevices is not a CCD image, but a series of electrophero-grams with a profile for each color scanned (nominallyBlue, Green, Yellow, and Red). Since these are difficult tointerpret the computer software provides decomposed sin-gle color graphs. Figure 8 contains an electropherogram ofthe single amplification Promega PowerPlex 16.2 Systemrun on an ABI PRISM 310 Genetic Analyzer.

One of the extremely useful characteristics of fluores-cent imaging devices is that the amount of light read by thedetection device is quantified. The electropherograms pro-duced by the real time scanners is quantitative. However,the CCD image can also be used as a scanning densitome-ter to determine the amount of light in each peak. Sincethere is one fluorescent molecule per band the amount offluorescence is linear with the number of molecules ina band. This allows for many different types of analysisto be performed on the data generated. One of the moreimportant of these is the ability to detect mixtures (seefollowing).


The manufacturers of the kits have done forensic vali-dations of the kits; however, each laboratory is responsi-ble for the individualized validation required before test-

ing. The guidelines for those validations for laboratoriesin the United States are governed by the DNA AdvisoryBoard Guidelines (as of October 1, 1998) as implementedby ASCLD-LAB. Other areas of the world are regulatedby other guidelines, unless they are also ASCLD-LABaccredited.

A. STR Detection

The major difference in the typing of the STR loci is theability to include an internal size standard if the detectiondevice used has multicolor capability. Under the TWG-DAM Guidelines forensic samples are to be placed ad-jacent to an allele ladder, as seen in Fig. 7 (PCR–STR).Since the Beckman Genomyx SC only has two filters (flu-oroscein and TMR) an internal ladder could not be used, sothe adjacent ladder format is used. In this situation there isno special preparation for detection. When the four-colorHitachi FMBIO II Fluorescent Scanner, ABI Prism 377or 310 is used, an internal standard is used to size theDNA fragements. As part of the electrophoresis setup aROX ladder is added to PE Biosystems amplified prod-ucts while a CRX ladder is added to Promega kits. (SeeFigure 8 for example.) Amplified products including the

FIGURE 8 Electropherogram of a single DNA sample amplifiedusing the 16-locus prototype PowerplexTM 16.2 System detectedwith the ABI PRISM® 310 Genetic Analyzer. All 16 loci wereamplified in a single reaction and detected in a single capillary.The fluorescein-labeled loci (D3S1358, THO1, D21S11, D18S51,and Penta E) are displayed in blue, the TMR labeled loci (Amel-ogenin, VWA03, D8S1179, TPOX, and FGA) are displayed inblack, and the loci labeled with a new dye (D5S818, D13S317,D7S820, D16S539, CSF1PO, and Penta D) are displayed ingreen. The fragments of the prototype ILS-500 size marker arelabeled with CXR and are shown in red. (Taken from Promega pro-motional literature.) (From Schanfield, M. S. (2000). Deoxyribonu-cleic Acid/Polymerase Chain Reaction-Short Tandem Repeats. In“Encyclopedia of Forensic Sciences” (Siegel, J. A., Saukko, P. J.,and Knupfer, G. C., eds.), Academic Press, London, p. 533.)

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alllelic size ladders. The internal size standard is used tosize all fragments within a lane by detector supplied soft-ware and assigns repeat numbers from the allele laddersizings.

B. The Use of Internal Size Standards

It was previously demonstrated that within gel variationin DNA migration could be compensated for by placing asize ladder within the lane and measuring each fragmentwith the internal size standard. This allows for highly pre-cise measurements of fragments. This is necessary sincethe electrophoresis systems used to detect the STR locimust have the capability of resolving differences betweenalleles as little as one base pair, to make sure that the frag-ment sizes can be accurately converted to repeat numbers.This would not be critical if all STR were regular. That isto say always four repeats for the tetra nucleotide STRs.However, this is not the case. Some STR loci have one ormore common alleles that differ by only a single base pair(THO1, FGA, and D21S11). An example of this is seenin Fig. 7B, Panel 1, third sample from the left, in the thirdlocus from the top.


One of the major problems in the analysis of forensic evi-dence is posed by samples containing biological materialfrom more than one source. The large number of discretealleles at multiple loci make STR multiplexes and excel-lent tool for identifying components of mixtures.


In the United States, the FBI has started releasing reportsindicating that biological material originated from a spe-cific source, much as fingerprint examiners have done formany years. The FBI has decided that if the populationfrequency exceeds 1 in 230 billion the sample is individu-alized. Other laboratories have chosen high thresold levelssuch as 1 in 500 billion. Whatever the level chosen it isestimated that the average power of exclusion for these 13CODIS loci exceeds 1 in a million billion, and though itis possible to obtain a frequency more common than thatrequired for individualization it will occur infrequently.






Adams, D. E., Presley, L. A., and Baumstark, A. L., et al. (1991). “De-oxyribonucleic Acid (DNA) analysis by restriction fragment lengthpolymorphism of blood and other body fluid stains subjected to con-tamination and environmental insults,” J. Forensic Sci. 36, 1284–1298.

Von Beroldingen, C. H., Blake, E. T., Higuchi, R., Sensabaugh, G., andEhrlich, H. (1989). “Application of PCR to the analysis of biologicalevidence. In “PCR Technology: Principles and Applications for DNAAmplification” (H. A. Ehrlich, ed.), Stockton Press, New York, pp.209–223.

Blake, E., Mihalovich, J., Higuchi, R., Walsh, P. S., and Ehrlich, H.(1992). “Polymerase chain reaction (PCR) amplification of humanleukocyte antigen (HLA)-DQ oligonucleotide typing on biologicalevidence samples: casework experience,” J. Forensic Sci. 37, 700–726.

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Dieffenbach, C. W., and Dveksler, G. S. (1993). “Setting up a PCRlaboratory,” PCR Methods Appl. 3, 2–7.

Fregeau, C. J., and Fourney, R. M. (1993). “DNA typing with fluores-cently tagged short tandem Repeats: A sensitive and accurate approachto human identification,” BioTechniques 15, 100–119.

Haugland, R. P. (1996). “Handbook of Fluorescent Probes and ResearchChemicals, 6th Edition,” Molecular Probes, Eugene, OR, pp. 144–156.

Jeffreys, A., Wilson, V., and Thein, S. L. (1985). “Hypervariable ‘min-isatellite’ regions in human DNA,” Nature 314, 67–73.

Kimpton, K. P., Gill, P., Walton, A., Urquhart, A., Millinan, E. A., andAdams, M. (1993). “Automated DNA profiling employing multiplexamplification of short tandem repeat loci,” PCR Methods Appl. 3, 13–22.

Laber, T. L., O’Connor, J. M., Iverson, J. T., Liberty, J. A., and Bergman,D. L. (1992). “Evaluation of four Deoxyribonucleic Acid (DNA) ex-traction protocols for DNA yield and variation in restriction fragmentlength polymorphism (RFLP) sizes under varying gel conditions,”J. Forensic Sci. 37, 404–424.

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Walsh, P. S., Metzger, D. A., and Higuchi, R. (1991). “Chelex® 100 asa medium for simple extraction of DNA for PCR based typing fromforensic material,” BioTechniques 10, 506–513.

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Gene Expression, Regulation ofGoran AkusjarviUppsala University

I. IntroductionII. Definition of a Transcription Unit

III. Regulation of Transcription in EukaryotesIV. Regulation of Transcription in ProkaryotesV. Posttranscriptional Regulation of Gene



Exon In eukaryotes, the part of the precursor-RNA thatreaches the cytoplasm as part of a mRNA, rRNA, ortRNA (see also intron).

Intron The part of the precursor-RNA that is removedduring RNA splicing, before the mature mRNA, rRNA,or tRNA is transported to the cytoplasm (see alsoexon).

Nucleosome The basic structural unit used to condenseDNA in a cell. The nucleosome consists of a disc-shaped core of histone proteins (H2A, H2B, H3, andH4) around which an approximately 146-base-pair seg-ment of DNA is wrapped.

Precursor-RNA The nuclear RNA transcript producedby transcription of the DNA. The precursor-RNA con-tains both exonic and intronic sequences. Introns areremoved by RNA splicing before the mature RNA istransported to the cytoplasm.

Promoter The DNA sequence element that determinesthe site for transcription inititation for an RNA poly-merase.

RNA splicing The nuclear process by which introns are

removed from the precursor-RNA before the maturemRNA, rRNA, or tRNA is transported to the cytoplasm.

U snRNP An RNA protein complex consisting of uridine-rich small nuclear RNAs (U snRNAs) complexed witha common set of proteins, the Sm proteins, and UsnRNA-specific proteins. The U1, U2, U4, U5, and U6snRNPs are involved in RNA splicing. Other U snRNPsserve other functions in the cell.

OUR GENES are stored in a stable DNA molecule thatis faithfully replicated and transmitted to new daughtercells. Each cell in our body contains the same geneticinformation. The development of complex multicellularorganisms such as humans with highly specialized organsand cell types then arises through a complex regulationof expression of the genetic material. Recent advances inwhole-genome sequencing have suggested that the differ-ence between a simple organism like a bacteria and a muchmore complex organism like a human may only result fromas little as a 5- to 10-fold difference in the number of genes.Thus, a prototypical bacterial genome encodes for 2000–4000 genes, whereas the recently completed sequencing of


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the human genome suggests a total gene number slightlymore than 20,000. Although this estimate may be too low,it appears unlikely, based on other measurements, that thenumber of genes in humans will exceed 50,000. At a first,and even a second, glance this small difference in the num-ber of genes makes it difficult to understand why humansand bacteria are so different from each other.

The past decade has seen an explosive increase in in-formation about regulation of gene expression. This re-view summarizes some of the general themes that haveemerged. It is focused on expression of protein-encodinggenes in higher eukaryotes. At appropriate places a com-parison with gene expression in prokaryotes is made in anattempt to highlight similarities and to show differencesthat might provide some answers to how mechanistic dif-ferences in the regulation of genes may provide at leastpart of the solution of to how a complex organism likehumans may have arisen without an enormous increase inthe number of genes compared to prokaryotes.


Expression of the genetic information has been summa-rized in the so-called central dogma, which postulates thatthe genetic information in a cell is transmitted from theDNA to an RNA intermediate to protein. A major differ-ence between simple and complex organisms is the exis-tence of a cell nucleus. Thus, prokaryotes, which includethe bacteria and the blue-green algae, do not have a nu-cleus, whereas eukaryotes, which include animals, plants,and fungi, have cells with a nucleus that encapsulates theDNA. The basic mechanisms to regulate gene expressionin eukaryotes and prokaryotes are very similar, althougheukaryotes generally use more sophisticated methods tosqueeze out more information from the DNA sequence. Inprokaryotes on–off switches of transcription appear to bethe key mechanism to control gene activity, although othermechanisms also contribute to the control of gene expres-sion: transcriptional attenuation, transcriptional termina-tions, and posttranscriptional effects. In eukaryotes similarmechanisms are in operation. However, a key differencebetween prokaryotes and eukaryotes is the extensive useof RNA processing to generate a mature mRNA. Thus,eukaryotic genes are encoded by discontinuous DNA seg-ments that require a posttranscriptional maturation to pro-duce a functional mRNA. As will be discussed later inthis review, the requirement for RNA splicing may bea key to the development of a highly differentiated or-ganism like humans. The general postulate that one genemakes one protein was derived from genetic studies ofbacteriophages and does not apply to higher eukaryotes.Because of alternative RNA processing events a large frac-

tion of eukaryotic genes encode for multiple proteins (seeSection V).


A transcription unit represents the combination of regu-latory and coding DNA sequences that together make upan expressible unit, whose expression leads to synthesisof a gene product that often is a protein but also may bean RNA molecule. In prokaryotes, proteins in a specificmetabolic pathway are often encoded by genes that areclustered and transcribed into one polycistronic mRNA.A polycistronic mRNA encodes for multiple proteins. Insuch mRNAs, ribosomes are recruited to internal transla-tional initiation sites through an interaction between the16S ribosomal RNA and the so-called Shine–Delgarno se-quence located immediately upstream of the translationalstart codon that is used to initiate protein synthesis.

In eukaryotes, in contrast, the primary transcrip-tion product is a precursor-RNA that undergoes severalposttranscriptional maturation steps before it is trans-ported to the cytoplasm and presented to the ribosomes.Thus, the 5′ end of the pre-mRNA is capped early aftertranscription initiation by addition of an inverted methy-lated guanosine nucleotide (the m7G-cap), the pre-mRNAis cleaved at its 3′ terminus, and an approximately 250-nucleotide poly(A) tail is added posttranscriptionally; fi-nally, the pre-mRNA is spliced to remove the interven-ing intron sequences, and thus form the spliced mRNAwhich is transported to the cytoplasm. These posttran-scriptional processing events give eukaryotes a unique,very important level to control gene expression (see Sec-tion V.). Furthermore, a eukaryotic mRNA usually is func-tionally monocistronic. This means that even if the mRNAencode for multiple open translational reading frames, theopen reading frame closest to the 5′ end of the mRNAis typically the only one translated into protein. This re-sults from the fact that the eukaryotic ribosome recognizesthe mRNA by binding to the modified 5′ end of a mRNA(recognizing the cap nucleotide), whereas prokaryotic ri-bosomes recognizes internal Shine–Delgarno sequencesin the polycistronic mRNA.

Transcription involves synthesis of an RNA chain thatis identical in sequence to one of the two complemen-tary DNA strands. DNA sequence elements upstream ofthe initiation site for transcription make up the promoterthat binds the RNA polymerase responsible for synthesisof the precursor-RNA. Transcription can be subdividedinto at least three stages: (1) initiation, which begins byRNA polymerase binding to the double-stranded DNAmolecule and incorporation of the first nucleotide(s); (2)

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elongation, during which the RNA polymerase, by a pro-cessive mechanism, moves along the DNA template ina 5′-to-3′ direction and extends the growing RNA chainby copying one nucleotide at a time; and (3) termination,where RNA synthesis ends and the RNA polymerase com-plex disassembles from the transcription unit.

Because of space limitation this review will mostlycover RNA synthesis and maturation of protein-encodingmessenger RNAs (mRNAs). However, similar mecha-nisms are used to regulate synthesis of other types of RNAmolecules.


Eukaryotic cells contain three DNA-dependent RNA poly-merases which are responsible for synthesis of specificRNA molecules in the cell. RNA polymerase I is respon-sible for the synthesis of ribosomal RNA, RNA poly-merase II is responsible for the synthesis of protein-codingmRNA and four small stable RNAs involved in splicing(U1, U2, U4, and U5), and RNA polymerase III is respon-sible for the synthesis of small RNAs such as transfer RNA(tRNA) and 5S RNA and a whole array of small RNAs in-cluding U6, which is involved in splicing. The three poly-merases are large enzymes consisting of approximately 15subunits each. Many subunits are shared among the differ-ent polymerases, whereas others are unique and determinethe promoter specificity during the transcription process.All three eukaryotic RNA polymerases contain core sub-units that show a great homology with the Escherichia coliRNA polymerase, suggesting that the basic mechanism ofRNA synthesis evolved early during evolution and is con-served.

As will become important later in this review, thelargest subunit of RNA polymerase II contains a 52-times-repeated stretch of seven amino acids at the carboxy-terminus. This heptapeptide repeat is refereed to as thecarboxy-terminal domain (CTD) and contains serines anda tyrosine that contribute to transcriptional regulation viareversible phosphorylation. The RNA polymerase that as-sembles at the promoter contains an unphosphorylatedCTD tail. This CTD tail anchors the polymerase to thepromoter by making interactions with TFIID bound atthe TATA box. The release of the RNA polymerase fromthe promoter (i.e., start of elongation) is associated with aphosphorylation of the CTD tail.

A. Structure of a Eukaryotic Promoter

The transcriptional activity of a prototypical RNA poly-merase II gene is regulated by a series of DNA sequence

FIGURE 1 A schematic model for preinitiation complex formationon a core promoter (a) and a promoter regulated by enhancer-binding transcription factors (b). The figure is meant to illustratethat the TATA-binding factor TBP is sufficient for basal transcrip-tion, whereas enhancer-dependent transcription requires TFIID,which consists of TBP plus the TBP associated factors (TAFs).Pol II, RNA polymerase II; GTFs, general transcription factors;INR, initiator region.

elements that can be subdivided into the core promoter ele-ment, which consists of the transcriptional start site and theTATA element, and upstream regulatory elements, whichare needed for regulated transcription (Fig. 1).

1. The Core Promoter

The TATA element located 25–30 base pairs (bp) upstreamof the transcription initiation site is critical for formationof the preinitiation complex by functioning as the bindingsite for the TATA-binding protein (TBP). The core pro-moter is sufficient to direct basal (unregulated) transcrip-tion. In some genes the transcriptional start site includesan initiator (Inr) element that binds specific factors thatmay substitute for the TATA box in recruiting the basaltranscriptional machinery to the promoter. Although thecore promoter is of fundamental importance for bindingof the general transcription apparatus, the composition ofelements may influence regulation of promoter activity.

2. Upstream Regulatory Factors

Upstream activating sequences (UAS), or transcriptionalenhancer elements, are binding sites for transcription fac-tors that stimulate RNA synthesis. The term UAS is used todescribe DNA sequence elements that are located close to

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the core promoter, so-called promoter proximal elements.UAS elements are typically located within 200 base pairsupstream of the transcription initiation site. Enhancer se-quences are DNA segments containing binding sites formultiple transcription factors that activate transcriptionindependent of their orientation and at a great distance[up to 85 kilobases (kb)] from the start site of transcrip-tion. Enhancer elements can be located either upstream ordownstream of the transcription initiation site. Enhancersequences activate transcription in a position-independentmanner because they become spatially positioned close tothe core promoter through bending of the DNA molecule(Fig. 1).

In addition to enhancer elements, eukaryotic promoterscontain upstream repressor elements, which block RNAsynthesis by various mechanisms by recruiting factors thatinterefere with enhancer factors or directly block RNApolymerase II recruitment. A third class of DNA sequenceelements regulating transcription are the transcriptionalsilencers. A classical silencer represses transcription ina position- and orientation-independent fashion. The si-lencer element is thought to block transcription by func-tioning as the nucleation site for binding of histones orsilencing proteins that coat the region, thereby making thepromoter inaccessible for RNA polymerase recruitment.

The human genome encodes for several thousand dif-ferent transcription factors. Promoters that contain com-binations of binding sites for different transcription fac-tors regulate different genes. Thus, for example, a genespecifically expressed in the liver or the brain uses liver-or brain-specific enhancer binding transcription factors,respectively, to achieve a tissue-specific gene expression.The basal transcriptional machinery appears to a large ex-tent to be the same in all cell types.

B. Regulation of Promoter Activity

From a regulatory point of view it is important to notethat TBP is sufficient to recruit RNA polymerase II anddirect basal transcription from the core promoter. How-ever, the basal transcription factor TFIID has been shownto play a central role in activated transcription by bindingto the TATA element in the core promoter and facilitat-ing the recruitment of the RNA polymerase holoenzymeto the promoter (Fig. 1). TFIID is a multiprotein complexconsisting of TBP and approximately 11 TBP-associatedfactors (TAFs). The TAFs have been shown to be essen-tial for regulated transcription by mediating contact withenhancer binding factors. Thus, TBP is sufficient for con-stitutive transcription but TAFs are necessary for regulatedtranscription (Fig. 1). In vitro studies suggest that assem-bly of an initiation-competent RNA polymerase at a pro-moter can be subdivided into several steps where different

basal transcription factors are sequentially recruited to thepromoter. However, fractionation experiments have shownthat on certain promoters the RNA polymerase and mostor all of the general transcription factors may be recruitedas a single complex. In vivo activation of the thousands ofpromoters present in the human genome may use a largespectrum of mechanistic possibilities.

An important finding was the observation that UAS-binding transcription factors are modular in structure witha DNA-binding domain and an effector domain that couldbe exchanged without losing their predicted biologicalactivity. The effector or activation domains in differenttranscription factors perform the same task but have dif-ferent properties, for example, consisting of acidic blobs,proline-rich, glutamine-rich, or serine/threonine-rich se-quences. Different classes of UAS-binding transcriptionfactors may transmit a signal to the basal promoter com-plex by making specific contacts with different TAFs. Forexample, an interaction between the UAS-binding tran-scription factor SP1 and TAF-110 has been shown tobe necessary for SP1-mediated activation of transcrip-tion. Collectively stabilized protein–protein interactionsbetween UAS-binding factors and the general transcrip-tional factor TFIID are likely to facilitate recruitment ofthe RNA polymerase to the core promoter element, and asa consequence increase the transcriptional activity of thepromoter (Fig. 1).

Transcription factors can be subdivided into familiesbased on the structural feature of the DNA-binding do-main. Thus, the DNA-binding domain may interact withthe DNA through structural types like the helix-turn-helixmotif found in homeodomain proteins, zinc fingers, orleucine-zipper-basic DNA-binding domain motifs. Het-erodimerization between members of UAS-activatingtranscription factors belonging to such structural types isnot uncommon and has been shown to increase the reper-toire by which transcription factors can interact with dif-ferent promoter sequences. For example, the prototypicalAP1 transcription factor, which belong to the leucine-zipper family of transcription factors, consists of a het-erodimer of c-jun and c-fos. It binds to its cognate DNAmotif with a higher affinity than, for example, a c-jun–c-jun homodimer or a JunB–c-fos heterodimer. The com-binatorial complexity is further increased by the fact thatc-jun may form heterodimers with members of the ATFfamily of transcription factors. Thus, heterodimerizationbetween different members of a transcription factor fam-ily is an important mechanism to generate factors withalternative DNA-binding specificity.

When the RNA polymerase leaves the promoter, TFIIDremains bound at the TATA element and is ready to help asecond RNA polymerase to bind and initiate transcriptionat the same promoter. The activity of TFIID appears also

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to be regulated by inhibitory proteins that interact withTBP. Such TBP–inhibitory protein complexes may servean important regulatory role by keeping genes which havebeen removed from inactive chromatin in a repressed butrapidly inducible state.

1. Regulation of Transcriptionby Chromatin Remodeling

During the last decade a wealth of information has demon-strated the significance of the chromatin template for thetranscriptional activity of a promoter. It has been knownfor decades that the DNA in our cells is wrapped around aprotein core called the nucleosome. The nucleosome con-sists of two copies each of four histones: H2A, H2B, H3,and H4. The DNA is packaged into either a loose struc-ture called euchromatin or a more highly ordered structurecalled heterochromatin. The heterochromatin fraction istranscriptionally inactive, whereas active genes are foundin the euchromatin fraction of the DNA.

Histones are modified by acetylation, phosphorylation,methylation, and ubiqutination. During recent years animpressive amount of work has demonstrated the sig-nificance of reversible histone acetylation as a regu-latory mechanism controlling gene expression. Severallysines on the amino-terminal tail of each core histone canbe acetylated. Lysines are negatively charged and makestrong interaction with the phosphate backbone of DNA,thereby preventing basal transcription factors like TBPfrom interacting with DNA. Acetylation of lysines neu-tralizes this negative charge and reduces the electrostaticinteraction of the histones with the DNA, thereby mak-ing the promoter region accessible for interaction withthe basal transcription machinery (Fig. 2). A considerableamount of work shows that a general theme in transcrip-tional regulation is that acetylation of core histones resultsin looser nucleosomal structure, which makes the DNAmore accessible for binding of transcription factors, andhence a gene more transcriptionally active. In contrast, hi-stone deacetylation has the opposite effect and functionsas a signal to repress transcription (Fig. 2).

Several transcriptional enhancer proteins have beenshown to activate transcription by binding so-calledcoactivator proteins which have histone acyltransferase(HAT) activity. The best characterized are Gcn5 (yeast),TAFII250, CBP, and p300. TAFII250, which is a compo-nent of the TATA-binding basal transcription factor TFIID,may activate transcription by inducing acetylation of hi-stones located in the vicinity of the TATA box. On theother hand, transcriptional repressor proteins have oftenbeen shown to inhibit RNA synthesis by recruiting histonedeacetyltransfereases (HDACs), which cause a condensa-tion of nucleosomes to a more compact, transcriptionally

FIGURE 2 Role of the nucleosome in gene expression. Recruit-ment of histone deacetylases (HDACs) to a promoter inhibits bind-ing of general transcription factors to the TATA element, therebyblocking transcription. Recruitment of histone acetylases (HATs)to the promoter results in acetylation of the amino-terminal tails ofthe core histones, thereby facilitating binding of the general tran-scription factors required for initiation of transcription. URS, up-stream repressor sequence; UAS, upstream activating sequence.

inactive structure (Fig. 2). In yeast, HATs and HDACsare found in multiprotein complexes such as SAGA andSin3 complexes, respectively. The equivalent, and addi-tional, multienzyme complexes are also found in highereukaryotes.

In addition, the nucleus contains so-called chromatinremodeling factors, such as the Swi/Snf complex, whichhas the capacity to reposition nucleosomes and transientlydissociate the DNA from the surface of the nucleosome.Depending on the promoter context, chromatin remod-eling factors may cause an activation or repression oftranscription.

It is likely that other histone modifications, such asphosphorylation, ubiquitinilation, and methylation, alsoplay a significant regulatory role in transcriptional con-trol of promoter activity, although the importance of thesemodifications has not yet been characterized to the sameextent as has that of reversible acetylation. The main con-clusion from these studies is that a linear assessment of the

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DNA sequence elements capable of binding transcrip-tional activator or repressor proteins only tells us partof the story, namely which factors have the capacity tocontrol promoter activity. However, actual RNA synthesisrequires a complex interplay between UAS-binding fac-tors and the chromatin or the chromatin remodeling factorsthat have positive or negative effects on promoter activity.

2. Regulation of Transcription Factor Activity

The activity of a UAS-binding transcription factor issubjected to a posttranslational regulation. There are inprinciple three ways that the activity of an UAS-bindingtranscription factor may be tuned (Fig. 3); covalent (likephosphorylation) or noncovalent (like hormone binding)modification of the UAS-binding factor, or variation ofthe subunit composition (like binding of an inhibitory pro-tein). These mechanisms may be used individually or incombination with other mechanisms to regulate transcrip-tion. To illustrate the flexibility of transcriptional con-trol in eukaryotic cells two examples are presented. Thefirst example concerns the activation of steroid hormone-dependent gene transcription (Fig. 4). Steroid hormonesare a group of substances derived from cholesterol whichexert a wide range of effects on processes such as growth,metabolism, and sexual differentiation. A prototypicalmember of a steroid hormone-inducible transcription fac-tor is the glucocorticoid receptor (GR). In the absence ofhormone this receptor is found as a monomer in the cy-

FIGURE 3 Three common mechanisms to regulate transcriptionfactor activity.

FIGURE 4 A schematic drawing showing the activation of gluco-corticoid receptor (GR) by steroid hormone binding, which resultsin the dissociation of the cytoplasmic GR–hsp90 complex, fol-lowed by GR dimerization and translocation of GR to the nucleus.

toplasm complexed to the heat-inducible hsp90 protein.Treatment with steroid hormones results in the releaseof GR from hsp90, which renders GR free to dimer-ize and move to the nucleus, where it binds to its cog-nate DNA sequence element and activates transcription(Fig. 4). In addition to inducing a dissociation of the re-ceptor from hsp90, ligand binding also induces a confor-mational change in the activation domain of GR, suchthat the activation domain binds transcriptional coacti-vator proteins that stimulate transcription. Interestingly,

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FIGURE 5 Activation of transcription of heat-shock genes inmammalian cells. The heat shock factor (HSF) is present as amonomer in normal cells. An increase in temperature results ina trimerization of HSF, which binds to the heat shock element(HSE). HSF is activated as a transcriptional enhancer protein byphosphorylation.

heat-shock proteins do not regulate other nuclear hormonereceptors, such as the retinoic acid and thyroid hormonereceptors. Thus, these receptors bind DNA in the absenceof the ligand. In this case ligand binding results in a con-formational change of the activation domain permittingbinding of coactivator proteins.

The second example concerns heat-shock activation oftranscription in mammalian cells (Fig. 5). When cells aresubjected to an elevated temperature (heat shock) they re-spond by activating synthesis of a small number of genesencoding for so-called heat-shock proteins. These pro-teins serve an important function during heat shock bybinding to cellular proteins, which become denatured bythe increase in temperature. Subsequently, the heat-shockproteins help to renature the proteins to their native con-formation. Transcription of heat-shock genes is controlledby the heat-shock transcription factor (HSF), which bindsto the heat-shock element (HSE) found in the promoterof all genes regulated by heat shock. HSF is activated bytwo mechanisms (Fig. 5). Thus, in normal cells HSF existsas a monomer. An increase in temperature results in un-folding of HSF, which exposes the DNA-binding domainand allows it to bind to other HSFs and form a trimer thatbinds to the HSE. However, binding of HSF to DNA isnot enough to activate transcription. Thus, HSF needs tobe modified by phosphorylation before it activates tran-scription of the heat-shock genes. Interestingly, TFIID isbound to the TATA element in heat-shock genes also inuninduced cells. Thus, binding of an active HSF to the

HSE is needed for recruitment of the RNA polymeraseto the heat-shock promoter. The binding of TFIID to theuninduced promoter may help heat-shock genes respondmore rapidly to an increase in temperature.

Cell type and differentiation-specific gene expression isoften regulated by the availability of specific transcriptionfactors. Genes that are expressed in specific organs containbinding sites for cell type-specific transcription factors.Thus, tissue-specific transcription is often regulated bythe precise arrangement of regulatory UAS motifs in thepromoter, the availability of the cognate transcription fac-tors, and the way these transcription factors influence theactivity of the promoter. Thus, for example, the liver andthe brain encode for respectively liver- and brain-specifictranscription factors that ensure a tissue-specific expres-sion gene expression. Since transcription factors are typ-ically dimeric proteins, the exact composition of the twopartners may vary among cell types and have differenttranscription regulatory properties.

3. Regulation of Transcription Elongation

Although transcription initiation has only been discussed,RNA polymerase elongation is also an important step inregulating gene expression in eukaryotes. Thus, there areseveral examples where the RNA polymerase halts at spe-cific pause sites during elongation. To be able to com-plete the synthesis of the precursor-RNA the polymerasehas to be able to override this attenuation of transcrip-tion. The best-characterized example is the human im-munodeficiency virus (HIV) Tat protein, which binds to astem-loop structure at the 5′ end of the HIV transcript, theTAR sequence. In the absence of the Tat protein, HIV tran-scription terminates approximaqtely 50 nucleotides down-stream of the initiation site. When Tat is present it bindsto the TAR sequence and recruits a cyclin T/Cdk9 com-plex which is responsible for phosphorylation of the CTDtail of RNA polymerase II, thereby alleviating termina-tion and permitting the RNA polymerase to synthesize thefull-length HIV genomic RNA.


A. Introduction

The mechanisms to initiate transcription in eukaryotes andprokaryotes are similar. As a comparison to control oftranscription in eukaryotes some key features in transcrip-tional control in bacteria will be given. Prokaryotic cellscontain only one type of RNA polymerase, which is re-sponsible for synthesis of all types of RNA: mRNA, rRNA,

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and tRNA. The core polymerase is a four-subunit enzymeconsisting of two a, one b, and one b′ subunit. However,the holoenzyme, which is the complete enzyme, containsthe core polymerase plus the sigma factor, which mayregarded as the prokaryotic equivalent of the general tran-scription factors found in eukaryotes. The sigma factor isrequired for proper RNA polymerase binding to a prokary-otic promoter. After initiation of transcription the sigmafactor leaves the polymerase complex and elongation istaken care of by the core polymerase.

Similar to a eukaryotic promoter a prototypical prokary-otic core promoter contains a conserved TATAAT locatedat position −10 relative to the transcription start site andresembling the eukaryotic TATA element. In addition,prokaryotic promoters contain a conserved TTGACA lo-cated at position −35. The spacing between the two ele-ments is of critical importance for the efficiency by whichthe RNA polymerase binds to the promoter. The exact se-quences at the −10 and −35 positions vary slightly fordifferent transcription units. Usually promoters that havea better homology to the consensus sequences also initiatetranscription more efficiently. An important mechanism toregulate the transcriptional activity of a prokaryotic pro-moter is to provide the core polymerase with differentsigma factors. Thus, different sigma factors determine pro-moter specificity by recognizing −10 and −35 elementswith different base sequences. This strategy mediates theheat shock response and the regulated expression of genesduring developmental processes. For example, sporulationin Bacillus subtilis uses a cascade of different sigma fac-tors to cause the transformation of a vegetative bacteriumto a spore.

The existence of transcriptional enhancers similar tothose found in eukaryotic cells has also been described inprokaryotes. For example, the enhancer-binding proteinnitrogen regulatory protein C (NTRC) in the glnA pro-moter from Salmonella typhimurium activates transcrip-tion from a distance by means of DNA looping. NTRCstimulates transcription by a transient contact between theactivator and the polymerase. This catalyzes an unwindingof the DNA at the promoter, which then allows the RNApolymerase to initiate transcription.

B. The Lac Operon

Transcriptional repression is a key mechanism to con-trol the activity of prokaryotic promoters. Enzymes usedin a specific metabolic pathway are often organized intoan operon that is transcribed into a single polycistronicmRNA. Specific repressor proteins then control the tran-scriptional activity of the operon by regulating RNA poly-merase binding to the promoter. Repressor proteins areDNA-binding proteins that typically block RNA poly-merase access to the −10 and/or −35 regions in the pro-

FIGURE 6 Regulation of the lac operon in E. coli. The lac I geneencodes for a transcriptional repressor protein that binds to anoperator sequence in the lac operon, thereby preventing synthesisof the structural genes required for metabolism of lactose. If E. coliis grown on lactose as the sole carbon source, lactose binds tothe lac I repressor protein and inactivates it as a repressor of lacoperon transcription. As a consequence, the β-galactosidase (lacZ ), the permease (lac Y ), and the β-galactosidase transacetylase(lac A) enzymes are synthesized.

moter or transcription elongation by associating with anoperator sequence that is positioned downstream of thestart site of transcription. Usually these regulatory pro-teins undergo allosteric changes in response to binding ofa specific ligand. The paradigm of a prokaryotic operonregulated by a specific repressor protein is the lac operonin E. coli. In this system synthesis of proteins necessaryfor usage of lactose as a carbon source is repressed by thelac repressor protein if cells have the possibility to useglucose for growth. Thus, in the presence of glucose thelac repressor binds to its operator sequence, which over-laps the transcription start site in the lac operon (Fig. 6),and blocks RNA polymerase binding to the lac promoter.If cells are grown on lactose as the carbon source, lac-tose functions as an inducer of lac operon transcription bybinding to the lac repressor and converting it to an inac-tive form that does not bind DNA (Fig. 6) and thereforeis unable to inhibit transcription of the lac operon. Thepolycistronic lac mRNA encodes for the specific proteinsnecessary for metabolism of lactose. The lac operon repre-sents an example of an inducible system where an induceractivates transcription. However, inducers can also havethe opposite effect and repress transcription of an operon,like the trp operon in E. coli.


Expression of eukaryotic genes is not only controlledat the level of initiation of RNA synthesis. Thus, the

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precursor-RNA synthesized by RNA polymerase II un-dergoes several posttranscriptional modifications before amature mRNA is formed. For example, the transcript iscapped at its 5′ end, the 3′ end is generated by a specificcleavage polyadenylation reaction, and intronic sequencesare removed by RNA splicing.

Early after initiation of transcription, when the nascentRNA chain is 25–30 nucleotides long, the 5′ end is modi-fied by addition of an inverted 7-methylguanosine, the capnucleotide. The capping enzymes are brought to the tran-scribing polymerase by specific association with the hy-perphosphorylated form of the CTD tail on RNA poly-merase II. As mentioned above, the CTD tail becomesphosphorylated when RNA polymerase II progress fromthe initiation to the elongation phase of RNA synthesis.Since RNA polymerases I and III do not have a CTD tail,only RNA polymerase II transcripts are capped. The capplays a crucial role in initiation of translation by bindingthe translational initiation factor eIF4F required for therecruitment of the small subunit of the ribosome to themRNA. The translational start site is then identified by ascanning mechanism where the ribosome usually selectsthe first AUG triplet as the start codon for protein synthe-sis. The selective addition of a cap to RNA polymeraseII transcripts therefore provides a logical explanation towhy this class of RNAs is used for translation. Polyadeny-lation and RNA splicing are key mechanisms to regulateeukaryotic gene expression and are therefore described inmore detail below.

A. Exons and Introns: General Considerations

Virtually all prokaryotic genes are encoded by a colli-near DNA sequence: the concept one gene, one mRNA.In contrast, most eukaryotic genes are discontinuous, withthe coding sequences (exons) interrupted by stretchesof noncoding sequences (introns). Introns are present atthe DNA level and in the primary transcription productof the gene (the precursor-RNA), and are removed byRNA splicing before the mature mRNA is transported tothe cytoplasm. Recent experiments suggest that splicingis necessary for efficient transport of intron-containingprecursor-RNAs. Introns have been found in all types ofeukaryotic RNA—mRNA, rRNA, and tRNA. Because ofspace limitation, only introns in protein-encoding geneswill be described.

The number of introns in mRNA-encoding genes variesconsiderably among genes. For example, c-jun, histone,heat-shock, and the α-interferon genes have no introns,whereas the gene for dystrophin has more than 70 introns.Also, the size of introns can vary from less than 100 nu-cleotides to several million nucleotides in length. The ex-treme example is the Drosophila Dhc7 gene, which con-tains a 3.6 million-nucleotide-long intron. This intron is

approximately double the size of most bacterial genomesand takes days to transcribe. In contrast, exons are typi-cally short, usually less than 350 nucleotides. This comesfrom the fact that splice sites used to define the borders ofthe splicing reaction are defined across the exon, not theintron—the so-called exon definition model (see below).Some eukaryotic genes are remarkably large. For exam-ple, the human gene for dystrophin covers approximately2.4 million base pairs. The RNA polymerase that initi-ates transcription requires approximately 20 hr to synthe-size the full-length precursor-RNA. Subsequently, morethan 99.5% of the transcript is removed by RNA splicing.Thus, the final mRNA that is transported to the cytoplasmis only around 14,000 nucleotides. The extreme lengthsof eukaryotic genes place a high demand on the stabilityof the transcription complex. Thus, an RNA polymerasethat binds to a promoter must stay attached for days withthe DNA template to be able to complete synthesis of thelongest genes.

It is interesting to note that introns in eukaryotic genesalmost always interrupt the protein-coding portion of theprecursor-RNA. Thus, introns are rarely found after thetranslational stop codon, within the 3′ noncoding portionof the mRNA. This organization is significant since thepresence of an intron downstream of the translational stopcodon in a reading frame is sensed as a signal that theprecursor-RNA has been incorrectly spliced or for otherreasons is defective, and will not produce the correct pro-tein after translation in the cytoplasm. Such nuclear tran-scripts are sent for destruction by a mechanism that iscollectively called the non-sense-mediated mRNA decaymechanism. How the translational reading frame is read al-ready in the nucleus is not known. The easiest explanationwould be that there exists a nuclear ribosome-like structurethat scans the spliced mRNA for a full-length translationalreading frame before the mRNA is transported to the cy-toplasm. However, this question is controversial and hasnot been proven.

1. Mechanism of RNA Splice Site Choice DuringSpliceosome Assembly

The sequence elements used to specify the splice sitesare remarkably short and degenerate in a eukaryoticprecursor-RNA. Thus, short conserved sequence motifsat the beginning (5′ end) and the end (3′ end) of the intronguide the assembly of a large RNA protein particle, thespliceosome (Fig. 7), which catalyzes the cleavage andligation reactions necessary to produce the mature cy-toplasmic mRNA. The nucleus of eukaryotic cells con-tains several abundant low-molecular-weight RNAs, so-called U snRNAs. The U snRNAs derive their namefrom the fact that they were initially characterized asRNAs rich in uridines. Five of these U snRNAs (U1,

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FIGURE 7 A simplistic model for spliceosome assembly. (a) The 5′ splice site and the branch site are defined via adirect base pairing between the RNA components of the U1 and U2 snRNPs and the precursor-RNA, respectively.(b) Efficient recruitment of the U snRNPs to the spliceosome is aided by non-snRNP proteins. Thus, SR proteinsfacilitate U1 snRNP binding to the 5′ splice site, whereas U2AF binds to the polypyrimidine tract at the 3′ splicesite and helps U2 snRNP binding to the branch site. The U4/U6–U5 triple snRNP is recruited to form the maturespliceosome. SR proteins bring the 5′ and 3′ splice sites in close proximity for the catalytic steps of splicing by makingsimultaneous contact with splicing factors binding to the 5′ and 3′ splice sites, respectively (see also Fig. 8).

U2, U4, U5, and U6), ranging in size from 107 to 210nucleotides, have been shown to participate in splicing.In vivo the snRNAs are found complexed to 6–10 pro-teins, generating the so-called small nuclear ribonucle-oprotein particles (snRNPs). Some snRNP proteins are

shared among different U snRNPs, whereas other snRNPproteins are unique to each U snRNP. During spliceosomeassembly the ends of the introns are in part identified byRNA–RNA base pairing between the precursor-RNA anda U snRNP (Fig. 7). For example, the 5′ splice site is

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recognized through a short base pairing between the U1snRNA and the precursor-RNA. Similarly, a base pairingbetween U2 snRNA and the branch point defines the 3′

splice site. Later during spliceosome formation the U5–U4/U6 triple snRNP is recruited. In the triple snRNP,U4 and U6 snRNP form an extensive base pairing. Thecatalytically active spliceosome is generated by confor-mational changes, which results in a breakage of the basepairing between U4 and U6 snRNP and formation of newU–U snRNA and U snRNA–precursor-RNA base pairings.It is generally believed, although not proven, that the UsnRNAs in the spliceosome are the enzymes that catalyzethe two transesterification reactions required to excise theintron.

2. Non-snRNP Proteins Required for Splicing

The spliceosome, which is a large RNA–protein complex,with a size similar to a cytoplasmic ribosome, also containsnumerous non-snRNP proteins which are important forcorrect splice site recognition. Assembly of the spliceo-some proceeds over several stable intermediates (Fig. 7).

Efficient recruitment of U2 and U1 snRNP to the 3′ and5′ splice sites also requires specific proteins. Here onlytwo factors will be described. The first is U2 snRNP aux-iliary factor (U2AF), which binds to the pyrimidine tractlocated between the branch site and the 3′ splice site inthe precursor-RNA. U2AF stabilizes U2 snRNP bindingto the branch site. The second factor is not one protein, buta family of proteins, designated SR proteins. SR proteinscontain one or two amino-terminal RNA-binding domainsand a carboxy-terminus rich in arginine (R) and serine (S)dipeptide repeats (the RS domain); hence the name SRproteins. Mechanistically, SR proteins appear to performthe same function in RNA splicing that transcriptionalenhancer proteins do in transcription initiation. Thus, SRproteins bind to splicing enhancer sequences through theirRNA-binding domains and stimulate spliceosome assem-bly by facilitating protein–protein interaction (Fig. 7).The RS domain functions as a protein interaction surfacethat makes contact with other SR proteins and so-calledSR-related proteins. Thus, many proteins involved in RNAsplicing contain RS domains. For example, SR proteins aidin efficient U1 snRNP binding to a 5′ splice site by inter-acting with the U1-70K protein, which is an RS-domaincontaining protein. However, in contrast to transcriptionalenhancer proteins, which active transcription irrespectiveof the position where they bind, SR protein function is po-sition dependent. Thus, in general, SR proteins functionas splicing-enhancer proteins if they bind to the exon andfunction as splicing-repressor proteins if they bind to theintron in the precursor-RNA.

The number of SR proteins found in mammalian cellsis surprisingly few considering the multitude of regulated

splicing events for which they are required. Thus, onlyaround 12 “true” SR proteins have been identified. Evenmore surprising, gene knockout experiments suggest thatonly one of the SR proteins is essential in Caenorhabditiselegans. Thus, disrupting the expression of the SR proteinASF/SF2 resulted in early embryonic lethality, whereasgene knockout of other SR proteins resulted in no changein phenotype. Probably, SR proteins show a large extentof functional redundancy, and disruption of one is com-pensated for by another SR protein. The essential role ofSR proteins in spliceosome assembly makes them primetargets for regulation of gene expression.

3. The Exon Definition Model

The conserved sequences at the 5′ and 3′ ends of the in-tron are surprisingly short considering the precision bywhich very large introns are excised during splicing. Theanswer to this puzzle appears to be resolved by the factthat the 5′ and 3′ splice sites that are joined in the splicingreaction are not recognized over the intron. Instead splicesites are recognized across the exons—the so-called exondefinition model. Thus, whereas introns can vary in lengthfrom less than 100 to more than 1 million nucleotides, internal exons in a precursor-RNA have a constant lengthand rarely exceed 350 nucleotides. The exon definitionmodel postulates that U2 snRNP binding to a 3′ splice sitemakes contact with U1 snRNP binding to the downstream5′ splice site (Fig. 8). If the 3′ and 5′ splice sites are too faraway the model postulates that the intervening sequenceis not recognized as an exon because U2 and U1 snRNPbinding to respective splice sites cannot interact with eachother. Once the exons have been defined in the precursor-RNA, adjacent exons are aligned for the splicing reaction.

B. Alternative RNA Splicing Is an ImportantMechanism to Generate Protein Diversity

A major difference in gene regulation between a prokary-otic and a eukaryotic cell is the existence of mechanismsin eukaryotic cells that permit one gene to express mul-tiple gene products. In bacteria a protein is encoded bya collinear DNA sequence. In contrast, in eukaryotes asingle gene may encode for thousands of proteins. Thus,the discontinuous arrangement of eukaryotic genes, withintrons interrupting the coding segments of the precursor-RNA, permit production of multiple, alternatively splicedmRNAs from a single gene. Examples of how a precursor-RNA can be alternatively spliced are shown in Fig. 9. This,of course, means that multiple proteins with different pri-mary amino acid sequence and biological activity can beproduced from a single eukaryotic gene. Of specific inter-est is that the production of alternatively spliced mRNAsin many cases is a regulated process, either in a temporal,

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FIGURE 8 The exon definition model. Exons in a precursor-RNAare recognized as units by U2 snRNP (U2) binding to the 3′ splicesite and U1 snRNP (U1) binding to the downstream 5′ splice site.Subsequently adjacent exons are defined across the intron. Inboth recognition steps SR proteins function as bridging proteins.

developmental, or tissue-specific manner. Changes insplicing have been shown to determine the ligand-bindingspecificity of growth factor receptors and cell adhesionmolecules and to alter the activation domains of transcrip-tion factors. For example, the fibronectin precursor-RNAis alternatively spliced in hepatocytes and fibroblasts. Infibroblasts two exons which are skipped in hepatocytes areincluded during the splicing reaction. These two exons en-code for protein domains that make fibroblast fibronectinadhere to many cell surface receptors. Fibronectin pro-duced in hepatocytes lacks these two exons and thereforeis translated to a hepatocyte-specific fibronectin proteinthat does not adhere to cells, allowing it to circulate in theserum.

The impact of alternative splicing on the coding capac-ity of a eukaryotic gene is mind-boggling. For example,

FIGURE 9 Examples of different patterns of alternative RNAsplicing.

the Drosophila DSCAM gene, which encodes for an axonguidance receptor, has been estimated to produce 38,016DSCAM protein isoforms by alternative splicing. This fig-ure is remarkable since the total gene number calculatedfrom the Drosophila DNA sequence suggests a total ofonly approximately 14,000 genes. Thus, a single Drosph-oila gene produces almost three times the number of pro-teins compared to the number of genes in Drosophila. TheDSCAM gene is not unique. There are many examples ofhuman genes, like those for neurexins, n-cadherins, andcalcium-activated potassium channels, that are known toproduce thousands of functionally divergent mRNAs. Alow estimate suggests that approximately 35% of all hu-man genes produce alternatively spliced mRNAs. Thus,the estimate of 20,000–50,000 genes in the human genomecould easily produce several hundred thousand, or million,proteins. Such differences in numbers are comforting be-cause they make it easier to explain how a complex organ-ism like humans with highly differentiated organs haveevolved without an enormous increase in the number ofgenes compared to bacteria.

1. Regulation of Alternative RNA Splicingby Changes in SR Protein Activity

With a few exceptions little is known about the mech-anistic details of how production of alternatively splicedmRNAs is regulated. However, it appears clear that the SRfamily of splicing factors partake in many regulated splic-ing events. SR proteins are highly phosphorylated, pri-marily within the RS domain. Thus, reversible RS domainphosphorylation has been shown to regulate SR protein

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FIGURE 10 Regulation of alternative RNA splicing by SR pro-tein phosphorylation. Hyperphosphorylated SR proteins presentin normal cells and early virus-infected cell bind to a specific re-pressor element in the adenovirus L1 precursor-RNA and blockspliceosome assembly at the IIIa 3′ splice site. This results in anexclusive production of the 52,55 K mRNA early after infection.Adenovirus induces a dephosphorylation of SR proteins late dur-ing infection, which alleviates the repressive effect of SR proteinson IIIa splicing, hence a shift to IIIa mRNA splicing.

interaction with other splicing factors and control alterna-tive RNA splicing.

One of the best-characterized examples is the humanadenovirus L1 unit (Fig. 10). The L1 unit produces twomRNAs, the 52,55K and the IIIa mRNAs, which are gener-ated by alternative 3′ splice site selection. Splicing duringan adenovirus infection is temporally regulated such thatthe IIIa mRNA is produced exclusively late during virusinfection. It has been shown that highly phosphorylatedSR proteins bind to an intronic repressor element and in-hibit IIIa splicing during the early phase of infection. Atlate times of infection IIIa splicing is activated by a virus-induced dephosphorylation of SR proteins. This changein the phosphorylated status of SR proteins reduces theirbinding capacity to the repressor element and hence re-sults in an alleviation of their repressive effect on IIIa 3′

splice site usage.

2. Maintenance of Sex in Drosophila:Sxl Regulation of Splicing

One of the most spectacular and best-characterized exam-ples where alternative RNA splice site choice is used toregulate gene expression is the somatic sex-determinationpathway in Drosophila melanogaster (Fig. 11). In this sys-tem sex determination has been shown to involve a cascadeof regulatory events taking place at the level of alternative

RNA splice site choice. The X chromosome encodes fortranscription factors that control Sex-lethal (Sxl) transcrip-tion. In females, which contain two X chromosomes, thedouble dose of these transcription factors results in an acti-vation of an early promoter of the Sxl gene. This promoteris inactive in males, which contain one X chromosome.The female-specific Sxl protein is an RNA-binding pro-tein that binds to certain pyrimidine tracts and outcom-petes U2AF binding to that site. Since the Sxl proteinlacks the splicing activator function of U2AF, Sxl bindingto a pyrimidine tract prohibits spliceosome formation atthe 3′ splice site. Thus, once made, the female-specificSxl protein autoregulates its own expression by ensuringthat exon 3 in the Sxl pre-mRNA is efficiently skipped,thereby establishing a female-specific splicing of the Sxlpre-mRNA (Fig. 11). In male flies exon 3 is incorporatedduring splicing, resulting in the translation of a function-ally inactive Sxl protein. This results from the fact that thethird exon in the Sxl pre-mRNA contains a translationalstop codon that causes a premature termination of transla-tion. In addition, Sxl controls the splicing of downstreamtargets in the sex determination pathway. Thus, Sxl regu-lates splicing of the transformer (Tra) precursor-RNA byrepressing usage of the male-specific 3′ splice site. Subse-qently the female-specific Tra protein complexes with theTra2 protein and activates a female-specific 3′ splice sitein the double-sex (Dsx) precursor-RNA (Fig. 11). In maleswhere a biologically inactive Sxl protein is expressed,the Sxl, Tra, and Dsx precursor-RNAs are processed bya default-splicing pathway, resulting in the developmentof male flies. The Dsx protein, the final protein in thecascade, is a transcription factor. The male- and female-specific Dsx proteins regulate development of flies alongthe male- or female-specific pathways.

It is widely accepted that alternative splicing is animportant mechanism to regulate gene expression dur-ing growth and development in eukaryotic cells. A largenumber of eukaryotic genes have been shown to maturealternatively spliced mRNAs, examples include growthfactors, growth factor receptors, intracellular messengers,transcription factors, oncogenes, and muscle proteins. Thenumber of examples is constantly increasing.

C. The Basic Mechanism of 3′ End Formation

1. Introduction

In eukaryotes all mRNAs except the histone mRNAs havea 200- to 250-long 3′ poly(A) tail. It is noteworthy thatthe 3′ end of a eukaryotic mRNA is not generated bytermination of transcription. Thus, the RNA polymerasecontinues to synthesize RNA beyond the actual 3′ endof the mature mRNA. Sequence analysis of a number ofRNA polymerase II genes has reveled two elements that

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FIGURE 11 The cascade of regulated alternative splicing events controlling Sxl, transformer, and double-sex ex-pression in Drosophila melanogaster. The positions of translational stop codons that will cause premature terminationof protein synthesis are indicated by Stop. See text for further details.

by biochemical assays have been shown to specify theposition of the 3′ end of a mRNA (Fig. 12). Thus, analmost invariable AAUAAA sequence located 25–30 nu-cleotides upstream of the cleavage site and a GU-rich se-quence located within 50 nucleotides downstream of thecleavage–polyadenylation site are critical for 3′ end for-mation. The AAUAAA sequence, which resembles theAT-rich TATA box important for transcription initiation,binds the essential cleavage–polyadenylation specificityfactor (CPSF). The GU-rich sequence binds the cleavage–stimulatory factor (CStF). In addition, two cleavage fac-tors, CFI and CFII, and the poly(A) polymerase (PAP)assemble to form an active enzyme complex that cleavesthe growing RNA chain and catalyzes the addition of the200- to 250-nucleotide poly(A) tail. The transcribing RNApolymerase terminates RNA synthesis in an ill-defined se-quence downstream of the cleavage–polyadenylation site.However, the cleavage–polyadenylation reaction has beenshown to be required for transcription termination, sug-gesting that breakage of the primary transcript is coupledto termination of transcription, possibly through the ac-tion of 5′ exonucleases that degrade the downstream RNAchain generated by the cleavage reaction. Mechanistically,

this may be analogous to Rho-dependent transcription ter-mination in bacteria.

2. Regulation of Gene Expression at the Levelof Alternative Poly(A) Site Usage

In addition to control of alternative RNA splicing, eukary-otic cells frequently use alternative poly(A) site usage tofurther increase the coding capacity of a gene. As de-scribed above, the 3′ end of a eukaryotic mRNA is gener-ated by an endonucleolytic cleavage of the primary tran-script rather than termination of transcription. This meansthat the RNA polymerase transcribing a gene may passby several potential poly(A) sites before terminating tran-scription. Thus, the selection of different poly(A) signalsin a precursor-RNA can be used to regulate gene expres-sion. For example, if the first poly(A) signal is ignored,a second poly(A) signal further downstream in a tran-scription unit may be used to incorporate novel exons intothe mRNA. As an example of this type of regulation, theproduction of secreted or membrane-bound immunoglob-ulin M (IgM) is described. When a hematopoietic stem celldifferentiates to a pre-B lymphocyte it produces IgM as an

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FIGURE 12 Model for cleavage and polyadenylation of aprecursor-RNA in mammalian cells. The cleavage and polyadeny-lation specificity factor (CPSF) binds to the conserved AAUAAAsequence located 10–35 nucleotides upstream of the poly(A) site.The cleavage-stimulatory factor (CStF) binds to the GU-rich ele-ment located downstream of the poly(A) site. Binding of two cleav-age factors (CF1 and CF II) and the poly(A) polymerase (PAP) thenstimulates cleavage of the precursor-RNA. The PAP synthesizesthe 200- to 250-nucleotide-long poly(A) tail, while the downstreamRNA fragment is rapidly degraded.

antibody anchored to the plasma membrane. After bind-ing to an antigen the lymphocyte undergoes differentia-tion and produces the same IgM molecule as a secreted

FIGURE 13 Alternative polyadenylation signals in the constant region of IgM yields heavy chains that are eithermembrane bound (pre-B cells) or secreted (plasma cells). Note that only the exon structure of the 3′ part of the IgMtranscription unit is shown. The upstream region, which encodes for the antigen-binding domain, is identical in thetwo forms of IgM.

protein. The shift from making a membrane-bound orsecreted antibody is regulated at the level of alternativepoly(A) site usage (Fig. 13). Thus, in the unstimulatedB cell the first poly(A) signal is ignored and the down-stream poly(A) signal is used as the default poly(A) site.The last two exons incorporated in the mRNA encode fora membrane-binding domain that anchors the antibodyto the plasma membrane. After stimulation the upstreampoly(A) signal is activated resulting in the processing ofa mRNA that is translated to the same antibody but lack-ing the membrane-binding domain. Hence this antibody issecreted.

Also, a new translational reading frame, encoding fora completely different protein, may be produced by alter-native poly(A) site usage. For example, production of cal-citonin, which occurs in thyroid cells, and the calcitonin-related protein, which is produced in the brain, is regulatedby tissue-specific poly(A) site usage.

D. Transcription and RNA ProcessingAre Coordinately Regulated Events

Recent experiments suggest that transcription and RNAprocessing are tightly coupled events. Thus, the RNApolymerase that assembles at the promoter has been shownto associate with factors required for polyadenylation andSR-related proteins that may partake in RNA splicing.The CTD tail of RNA polymerase II appears to func-tion as a platform that recruits the RNA processing fac-tors to the preinitiation complex. Therefore, the elongat-ing RNA polymerase appears to have been loaded withthe factors required for polyadenylation, and probably

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FIGURE 14 Editing of the Apo-B mRNA in intestinal cells. A CAA codon is edited to a UAA translational stop signalresulting in the production of a shorter protein (Apo-B48) corresponding to the amino-terminal half of the Apo-B100protein expressed in liver cells.

deposits them at the polyadenylation signal used for 3′

end formation. Even more surprising, evidence has beenpresented suggesting that the composition of a promotermay specify alternative RNA splicing. Thus, the samegene under the transcriptional control of different pro-moters produces different types of alternatively splicedmRNAs. This finding suggests that enhancer binding tran-scription factors, in addition to stimulating recruitment ofthe RNA polymerase to the promoter, also may partakein the recruitment of selective RNA processing factors tothe CTD tail. The future will tell whether alternative RNAsplicing is regulated already at the level of transcriptioninitiation.

E. Other Mechanisms of PosttranscriptionalRegulation

Although this review has focused on control of gene ex-pression at the level of synthesis and processing of mRNA,there are additional mechanisms that make significant con-tributions to gene expression. For example, a few genesin vertebrates have been shown to use RNA editing toproduce different protein isoforms. The serum proteinapolipoprotein B (Apo-B) is expressed in two forms. TheApo-B100 is expressed in hepatocytes, whereas a shorterpolypeptide, Apo-B48, is expressed in intestinal epithelialcells. The cell-type-specific expression of Apo-B resultsfrom a posttranscriptional editing of the Apo-B mRNA inintestinal epithelial cells. Thus, a CAA codon encodingfor the amino acid glutamate is converted to a UAA stopcodon by cytosine deamination. As a result the Apo-B48protein translated from the edited mRNA in the intestinediffers from Apo-B100 by lacking the carboxy-terminus(Fig. 14). Both proteins bind to lipids. However, only theliver-specific Apo-B100 contains the carboxy-terminaldomain required for binding to the low-density lipopro-tein receptor, necessary for delivery of cholesterol to bodytissues.

In nuclear genes RNA editing appears to be rare. Also,editing in such genes is restricted to modification of singlenucleotides. In contrast, RNA editing in genes expressedin the mitochondria of protozoa, plants, and chloroplastsresults in a more dramatic change of the mRNA sequence.Thus, a precursor-RNA may be edited such that more than50% of the sequence in the mature mRNA is altered com-pared to the primary transcription product.

Gene expression is also regulated at other levels, such asnuclear to cytoplasmic transport of mRNA, translationalefficiency of mRNA, RNA and protein stability, or proteinmodification. As is the case for transcriptional regulation,control of gene expression at the level of translation oftenoccurs at the initiation step of the decoding process. Thus,not all mRNAs that reach the cytoplasm are used directlyto synthesize protein. In fact, as much as 10% of genes in aeukaryotic cell may be regulated at the level of translation.






Lodish, H., Berk, A., Zipursky, S. L., Matsudaria, P., Baltimore, D.,and Darnell, J. (2000). “Molecular Cell Biology,” Freeman andNew York.

Latchman, D. S. (1998). “Eukaryotic Transcription Factors,” AcademicPress, San Diego, CA.

Gesteland, R. F., Cech, T. R., and Atkins, J. F. (1999). “The RNA World,”2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor,New York.

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Immunology—AutoimmunityK. Michael PollardEng M. TanScripps Research Institute

I. Autoimmunity and AutoantibodiesII. Detection of Autoantibodies and Autoantigens

III. Perspectives


Antibody A protein product of a specific type of lym-phoid cell (B cell) which combines with a specificmolecular target, called an antigen.

Antigen A substance, usually protein, that combines withan antibody.

Autoantigen A constituent of the body that is recognizedby its immune system. An antibody that recognizes anautoantigen is called an autoantibody.

Chromophore A chemical group capable of selectivelight absorption resulting in the coloration of certainorganic compounds.

Neoantigen A new or different antigen.

IN NORMAL circumstances a healthy immune system istolerant of the tissue constituents of the host but intolerantof nonhost (or “nonself ”) matter such as invading viruses,bacteria, or other pathogens. In autoimmunity the immunesystem mounts responses against tissue constituents of thehost organism. This response is most commonly referredto as recognition of “self,” as opposed to “nonself.” Theself/nonself paradigm is argued by many to be the ba-sis for understanding how the immune system determines

whether or not to respond to a suspected challenge. Con-siderable debate has occurred regarding the definition ofself and non-self, with both “infection” and “danger” be-ing suggested as possible discriminators. Irrespective ofthe final outcome of these discussions, it is clear that idio-pathic autoimmunity constitutes a significant disruption inthe mechanisms that regulate the immune systems’ abil-ity to discriminate at the molecular level. Autoimmunitycan lead to devastating diseases such as systemic lupuserythematosus (SLE) and insulin-dependent diabetes mel-litus (IDDM). These two clinical syndromes reflect thetwo major types of autoimmunity. SLE is representativeof multisystem autoimmunity, in which disease processesmay involve a number of organs of the body. IDDM is il-lustrative of organ-specific autoimmunity, which is morerestrictive, targeting perhaps a single organ such as thepancreas. Advances in our understanding of the relation-ships between autoimmune diseases and the autoantibod-ies and autoantigens that characterize them have comeabout through the use of an ever-evolving armamentariumof techniques based in part on the physical sciences. Thischapter is an attempt to describe the history, principles,and methodology of those techniques that have proventhe most useful in the characterization of autoantibod-ies and autoantigens in both basic and applied research


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680 Immunology—Autoimmunity

arenas. To appreciate fully the significant roles these tech-niques have played it is necessary first to review brieflysome of the fundamental features of autoimmunity andautoantibodies.


Autoimmune responses constitute an attack by the im-mune system on the host. The responsible effector mech-anisms appear to be no different from those used to combatexogenous agents and include soluble products such as an-tibodies (humoral immunity), as well as direct cell-to-cellinteraction leading to specific cell-lysis (cell-mediated im-munity). Autoantibodies are therefore defined as antibod-ies produced by the host which recognize cellular or tissueconstituents of the host. No single mechanism has beendescribed that can account for the diversity of autoimmuneresponses or the production of autoantibodies. Perhaps themost perplexing and challenging aspect of autoimmunityand autoantibody elicitation is the identification of theevents involved in the initiation of the response. Althoughthese early events are poorly understood for most autoim-mune diseases, it is thought that an exogenous trigger mayprovide the first step in the initiation of some autoimmuneresponses. It is also uncertain how T and B cells, with re-ceptors for autoantigen, emerge from primary lymphoidtissues, having escaped the regulatory mechanisms thatnormally delete them or keep them in check, and maketheir way to secondary lymphoid tissues where they canbe activated to respond in an inappropriate manner. Stud-ies involving transgenic mice expressing neoautoantigenssuggest that possible mechanisms for emergence of au-toractive cells include avoidance of apoptotic elimination,escape from tolerance induction, and reversal of an anergicstate. Molecular identification of autoantigens, their pres-ence in macromolecular complexes, the occurrence of au-toantibodies to different components of the same complex,and the appearance of somatic mutations in the variableregions of autoantibodies have suggested that autoantigendrives the autoimmune response. These findings supportthe argument that activation of autoreactive cells occursin secondary lymphoid tissues. It remains unclear howautoantigens, particularly intracellular autoantigens, aremade available to the immune system and what molec-ular forms of these complex macromolecular structuresinteract with autoreactive lymphoid cells.

An important component in humoral autoimmune re-sponses is the appearance of autoantibody-secreting Bcells. The antibody secreted by a B cell is directed againsta single region (or epitope) on an antigen. An autoanti-body response can target a number of epitopes on any oneantigen, clearly showing that multiple autoreactive B-cell

clones are activated during an autoimmune response. Inthe systemic autoimmune diseases many autoantigens arecomplexes of nucleic acid and/or protein, and an autoim-mune response may target several of the components ofa complex. It is unknown whether the autoantibody re-sponses to the components of a complex arise simul-taneously, sequentially, independently, or through someinterrelated mechanism. Immunization studies in micehave suggested that some autoantibody responses mayarise sequentially, a process termed epitope spreading.Whether this mechanism is applicable to humans is underinvestigation.

In only a few instances have autoantibodies beenshown to be the causative agents of pathogenesis (e.g.,anti-acetylcholine receptor autoantibodies in myasthe-nis gravis, anti-thyroid stimulating hormone receptor au-toantibodies in Graves’ disease). It is noteworthy thatthese diseases are usually organ specific and that theirautoantigens are extracellular or on the surface of cellmembranes and therefore easily targeted by the immunesystem. In the non-organ-specific autoimmune diseasesystemic lupus erythematosus (SLE) anti-double-strandedDNA (dsDNA) autoantibodies have been shown to partic-ipate in pathogenic events by way of complexing withtheir cognate antigen to cause immune complex medi-ated inflammation. These examples show that in bothorgan-specific and systemic autoimmune diseases, in vivodeposition of autoantibody in tissues and organs hasclinical significance, as it indicates sites of inflamma-tion and possible pathological lesions. Detection of au-toantibody/autoantigen deposits in organ-specific autoim-mune diseases has particular significance because passiveinfusion of some organ-specific autoantibodies has beenfound directly to mediate pathological sequelea. In mostautoimmune diseases, however, it has not been deter-mined whether autoantibodies cause or contribute in anyway to disease. It is possible that autoantibodies are anindicator of the primary event or a secondary conse-quence of the underlying clinical condition; autoantibod-ies have been described as potential “reporters” of diseasemechanisms.

Diseases associated with autoantibodies can be dividedinto two broad groups: multisystem autoimmune diseases,in which autoantibodies react with common cellular com-ponents that appear to bear little relationship to the clin-ical syndrome, and organ-specific autoimmune diseases,in which autoantibodies have the ability to react with au-toantigens from a particular organ or tissue. In both sit-uations the specificity of the autoantibody can serve as adiagnostic marker (Table I). There are several features ofthe relationship between autoantibody specificity and di-agnostic significance within the multisystem autoimmunediseases that bear consideration. Autoantigens in these

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TABLE I Examples of the Clinical Diagnostic Specificity of Autoantibodiesa

Autoantibody specificitya Molecular specificity Clinical association

Organ-specific autoimmune diseases

Anti-acetylcholine receptor∗ Acetylcholine receptor Myasthenia gravis

Anti-TSh receptor∗ TSH receptor Graves’ disease

Antithyroglobulin∗ Thyroglobulin Chronic thyroiditis

Anti-thyroid peroxidase∗ Thyroid peroxidase Chronic thyroiditis

Antimitochondria∗ Pyruvate dehydrogenase complex Primary biliary cirrhosis

Antikeratinoctye∗ Desmoplakin I homologue Bullous pemphigoid

Antikeratinoctye∗ Desmoglien Pemphigus foliaceus

Multisystem autoimmune diseases

Anti-double-stranded DNA∗ B form of DNA SLE

Anti-Sm∗ B, B′, D, and E proteins of U1, U2, and U4–U6 snRNP SLE

Anti-nRNP 70-kDa, A, and C proteins of U1-snRNP MCTD, SLE

Anti-SS-A/Ro 60- and 52-kDa proteins associated with the hY1–Y5 SS, neonatal lupus, SLERNP complex

Anti-SS-B/La 47-kDa phosphoprotein complexed with RNA SS, neonatal lupus, SLEpolymerase III transcripts

Anti-Jo-1∗ Histidyl tRNA synthetase Polymyositis

Antifibrillarin∗ 34-kDa protein of box C/D containing snoRNP Scleroderma(U3, U8, etc.)

Anti-RNA polymerase 1∗ Subunits of RNA polymerase 1 complex Scleroderma

Anti-DNA topoisomerase 1 (anti-Scl-70)∗ 100-kDa DNA topoisomerase I Scleroderma

Anticentromere∗ Centromeric proteins CENP-A, -B, and -C CREST (limited scleroderma)

cANCA Serine proteinase (proteinase 3) Wegener’s vasculitis

a SLE, systemic lupus erythematosus; MCTD, mixed connective tissue disease; SS, Sjogren’s syndrome; cANCA, cytoplasmic antineutrophilcytoplasmic antibody; TSH, thyroid-stimulating hormone; CREST, calcinosis, Raynaud’s phenomenon, esophageal dysmotility, sclerodactyly,telangiectasia. Disease-specific diagnostic marker antibodies indicated by an asterisk.

diseases are components of macromolecular structuressuch as the nucleosome of chromatin and the small nuclearribonucleoprotein (snRNP) particles of the spliceosome,among others. Autoantibodies to different components ofthe same macromolecular complex can be diagnostic fordifferent clinical disorders. For example, the core pro-teins B, B′, D, and E, which are components of the U1,U2, and U4–U6 snRNPs and are antigenic targets in theanti-Smith antigen (Sm) response in SLE, are differentfrom the U1 snRNP specific proteins of 70 kDa, A, andC, which are targets of the anti-nuclear RNP (nRNP) re-sponse in mixed connective tissue disease (MCTD; seeTable I). It has also been shown that particular autoanti-body responses are consistently associated with one an-other. The anti-Sm response, which is diagnostic of SLE,is commonly associated with the anti-nRNP response, butwhen the anti-nRNP response occurs in the absence of theanti-Sm response it can support the diagnosis of MCTD.These two observations suggest that the snRNP com-plexes responsible for the autoantibody response againstthe spliceosome in MCTD may differ from the snRNPcomplexes that produce the antispliceosome response in

SLE. Other autoantibody responses demonstrate similarassociations and restrictions. The anti-SS-A/Ro response(see Table I) frequently occurs alone in SLE but the anti-SS-B/La response in Sjogren’s syndrome is almost alwaysassociated with the anti-SS-A/Ro response. Similarily theantichromatin response occurs alone in drug-induced lu-pus but is often associated with the anti-dsDNA responsein idiopathic SLE. Autoantibody specificities may occurat different frequencies in a variety of diseases, and theresulting profile consisting of distinct groups of autoan-tibodies in different diseases can have diagnostic value.In some cases the grouping of autoantibody specificities,such as the preponderance of antinucleolar autoantibod-ies in scleroderma (Table I), provides intriguing but asyet little understood relationships with clinical diagnosis.Unlike SLE, in which a single patient may have multipleautoantibody specificities to a number of unrelated nuclearautoantigens (e.g., DNA, Sm, SS-A/Ro), scleroderma pa-tients are less likely to have multiple autoantibody speci-ficities to nucleolar autoantigens that are unrelated at themacromolecular level (i.e., not part of the same macro-molecular complex).

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TABLE II Examples of Subcellular Structures and Domains Recognized by Autoantibodiesa

Autoantibody Molecular specificity Subcellular structure

Nuclear components

Antichromatin Nucleosomal and subnucleosomal Chromatincomplexes of histones and DNA

Anti-nuclear pore 210-kDa glycoprotein (gp210) Nuclear pore

Antilamin Nuclear lamins A, B, C Nuclear lamina

Anticentromere Centromere proteins (CENP) A, B, C, F Centromere

Anti-p80 coilin p80-coilin (80-kDa protein) Coiled or cajal body

Anti-PIKA p23- to 25-kDa proteins Polymorphic interphase kayrosomalassociation (PIKA)

Anti-NuMA 238-kDa protein Mitotic spindle apparatus

Nucleolar components

Antifibrillarin 34-kDa fibrillarin Dense fibrillar component of nucleolus

Anti-RNA polymerase 1 RNA polymerase 1 Fibrillari center of nucleolus

Anti-Pm-Scl 75- and 100-kDa proteins of the Granular component of nucleolusPm–Scl complex

Anti-NOR 90 90-kDa doublet of (human) Nucleolar organizer region (NOR)upstream binding factor (hUBF)

Cytosolic components

Antimitochondria Pyruvate dehydrogenase complex Mitochondria

Antiribosome Ribosomal P proteins (P0, P1, P2) Ribosomes

Anti-Golgi 95- and 160-kDa golgins Golgi apparatus

Antiendosome 180-kDa protein Early endosomes

Antimicrosomal Cytochrome P450 superfamily Microsomes

cANCA Serine proteinase (proteinase 3) Lysosomes

Antimidbody 38-kDa protein Midbody

Anti-centrosome/centriole Pericentrin (48 kDa) Centrosome/centriole

a NuMA, nuclear mitotic apparatus; Pm–Scl, polymyositis–scleroderma; cANCA, cytoplasmic antineutrophil cytoplasmicantibody.

The molecular spectrum of autoantigenic targets (seeTables I and II) together with their exquisite antigenicspecificity has made autoantibodies valuable reagents inmolecular and cellular biology. The most visually impres-sive demonstration of the usefulness of autoantibodies asbiological probes is the indirect immunofluorescence (IIF)test. Using this technique (see Section II.A), an increas-ing number of autoantibody specificities are being iden-tified that recognize cellular substructures and domains(Table II and Fig. 1). Autoantibodies against chromatinand DNA can be used to identify the cell nucleus. Othernuclear structures such as the nuclear lamina, which un-derlies the nuclear envelope, can be identified by anti-lamin autoantibodies as a ring-like fluorescence aroundthe nucleus (Fig. 1a). The nucleolus and its subdomainscan be identified by a variety of autoantibodies (Table II).Antifibrillarin autoantibodies, which recognize the highlyconserved 34-kDa fibrillarin [a component of some smallnucleolar RNP (snoRNP) particles], identify the dense fib-rillar component. Autoantibodies to RNA polymerase I,

although rare, have proven to be a useful marker for thefibrillar center of the nucleolus. A growing appreciationof the antigenic diversity of cellular constituents, togetherwith improvements in fluorescent microscopy, has led toidentification of autoantibodies reacting with a variety ofsubnuclear domains and compartments, some consider-ably smaller than the nucleolus. The coiled body, a smallcircular subnuclear structure originally described by theSpanish cytologist Santiago Ramon y Cajal in 1903, andnow named the Cajal body, is an example. Cajal bod-ies can be identified using autoantibodies that react withp80 coilin (Fig. 1d), an 80-kDa protein highly enrichedin Cajal bodies. Using other autoantibodies in colocaliza-tion studies, it has been found that Cajal bodies containother proteins, including fibrillarin (previously thoughtto be restricted to the nucleolus and prenucleolar bod-ies). Autoantibodies have also been identified that reactwith subcellular structures other than the nucleus (Fig. 1).Prior knowledge of the existence and relative distribu-tions of these subcellular organelles was instrumental in

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FIGURE 1 Immunofluorescence patterns produced by autoantibodies recognizing structural domains within the nu-cleus (a–f) and cytosol (g–l) of the cell. (a) Antinuclear lamin B1 antibodies identify the periphery of the nucleus;arrowheads show the reformation of the nuclear envelope during late telophase. (b) Anti-Sm antibodies localize theU1, U2, and U4–U6 snRNP particles as a speckled pattern, but are absent from metaphase cells (arrowhead). (c) Anti-PCNA antibodies recognize the auxiliary protein of DNA polymerase delta during active DNA synthesis, producingdifferent fluorescence patterns as cells progress through mitosis. (d) Anti-p80 coilin antibodies highlight subnucleardomains known as cajal or coiled bodies, which disappear during metaphase (arrowhead). (e) Antifibrillarin antibodiestarget the nucleolus, produce a characteristic clumpy pattern in interphase cells, and decorate the chromosomes fromlate metaphase until cell division (arrowheads). (f) Antibodies to centromeric proteins A, B, and C produce a discreetspeckling of the interphase nucleus and identify the centromeric region of the dividing chromosomes during cell divi-sion (arrowheads). (g) Anti-mitotic spindle apparatus antibodies identify spindle poles and spindle fibers during celldivision. (h) Antimidbody antibodies react with the bridge-like midbody that connects daughter cells following chromo-some segregation but before cell separation. (i) Anti-Golgi complex antibodies decorate the Golgi apparatus, whichin most cells is shown as a discreet accumulation of fluorescence in the cytoplasm. (j) Antimitochondrial antibodiesdemonstrate the presence of mitochondria throughout the cytoplasm; the discreet nuclear dots represent an addi-tional autoantibody specificity in this serum unrelated to mitochondria. (k) Antiribosome antibodies produce a diffusecytoplasmic staining pattern that spares the nucleus but may show weak nucleolar fluorescence, (l) Anticytoskeletalantibodies react with a variety of cytoskeletal components; in this case the antibody reacts with nonmuscle myosin.Original magnification: a–f and h–l, 350×; g, 700×.

identifying the structures recognized by these autoanti-bodies. Conversely autoantibodies, by virtue of their re-activity with individual autoantigens, have allowed celland molecular biologists insight into the molecular con-stituents of these same subcellular organelles.

Autoantibodies can be used to study changes in the size,shape, and distribution of subcellular structures during thecell cycle, viral infection, mitogenesis, or any cellularresponse that results in alterations of subcellular con-stituents. For example, anti-lamin B1 autoantiodies can be

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used to demonstrate the reformation of the nuclear lam-ina during telophase (Fig. 1a). Autoantibodies have alsoidentified unexpected distributions of autoantigens, suchas the distribution of the nucleolar protein fibrillarin tothe outer surface of the chromosomes during cell division(Fig. 1e, arrowheads). The localization of some autoanti-gens during the cell cycle has aided in their identification.Detection of proliferating cell nuclear antigen (PCNA)in S-phase cells (Fig. 1c) suggested its involvement inDNA synthesis, while the distribution of speckles alongthe metaphase plate produced by other antibodies (Fig. 1f,arrowheads) was a significant contribution to their identi-fication as autoantibodies against the centromeric proteinsA, B, and C.

A feature of autoantibodies that underscores theiruniqueness is their ability to recognize their target antigennot only from the host but also from a variety of species.The extent of this species cross-reactivity is dependenton the evolutionary conservation of the autoantigen andis related to the conservation of protein sequence. Oneexample is the snoRNP protein fibrillarin. Using autoanti-bodies in a variety of techniques, this protein can be foundin species as diverse as humans and the unicellular yeastSaccharomyces cerevisiae. cDNA cloning of fibrillarin hasconfirmed the expected high degree of conservation of theprotein sequence.

Autoantibodies react with the conserved sequence andconformational elements of their cognate antigens; thesefeatures have made them useful regents in the cloning ofcDNAs of expressed proteins from cDNA libraries froma variety of species. However, because of their reactivitywith the human protein, they have been used primarilyto clone the cDNAs and characterize the primary struc-tures of numerous human cellular proteins. The diversityof the targets that have been exploited by this approach isillustrated in Tables I and II.

Elucidation of the structure of the autoantigens that arethe targets of autoantibodies from systemic autoimmunediseases has revealed that many are functional macro-molecular complexes involved in nucleic acid or proteinsynthesis. A distinguishing feature of many of these com-plexes of nucleic acid and/or protein is that autoantibodiesdo not recognize all the components of the complex. Anextreme, but useful, example is the ribosome, which in eu-karyotes may contain more than 70 proteins. However fewof these proteins are recognized by autoantibodies, the ma-jor targets being the sequence-related P proteins (P0, P1,P2). Nonetheless, the use of autoantibodies that identifyspecific components of such complexes has aided in iden-tifying other subunits of these complexes, with profoundconsequences. Thus the initial identification of anti-Smand anti-nRNP autoantibodies in SLE led to the observa-tion that they recognize some of the protein components

of the snRNP particles, fueling subsequent studies thatshowed the snRNPs as components of the spliceosomecomplex that functions in pre-mRNA splicing.

As the molecular and functional associations of au-toantigens have become known, attempts to uncover theparticular role of individual autoantigens have revealedthat autoantibodies can directly inhibit the function oftheir cognate autoantigen. Although it remains to be de-termined, it seems likely that such inhibition reflects theinvolvement of conserved protein sequence or structurein functional activity. An increasing number of autoanti-bodies, many of unknown molecular specificity, recognizetheir autoantigen only in a particular functional state orphase of the cell cycle. Of the several examples known, thebest characterized is PCNA, which is the auxillary proteinof DNA polymerase delta and is recognized by autoanti-bodies only during mitosis, even though PCNA is presentthroughout the cell cycle. When a population of cells atdifferent stages of the cell cycle is used in immunofluo-rescence anti-PCNA, autoantibodies produce varying de-grees of fluorescence intensity, being negative for G0 cellsand highly positive for S-phase cells (Fig. 1c). Theseintriguing features of some autoantibodies have addednew dimensions to their biological usefulness and havesuggested that functionally active macromolecular com-plexes may play a role in the elucidation of autoantibodyresponses.


The most commonly used methods for the detection andcharacterization of autoantibodies, whether in clinicalmedicine or molecular/cellular biology, fall under sev-eral broad biophysical areas and include fluorescent, en-zymatic, and radiographic techniques. As described belowsome of these techniques have been specifically developedfor antibody detection, while others have been borrowedand/or adapted from the biological/biophysical sciences.The methodology employed in these techniques has beendescribed in limited detail to afford the reader the op-portunity of understanding how the technique is put intopractice at the laboratory bench, particularly in the studyof autoimmunity.

A. Immunofluorescence

1. History

Characterization of the antigenic specificity of serum an-tibody by immunofluorescent methods dates to the early1950s. In what was the forerunner of current technology,Weller and Coons, in 1954, used human serum to identify

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viral antigens in tissue culture monolayers infected withvaricella and herpes zoster. The human antibody bound toviral antigen in the infected cells was detected by a flu-orochrome covalently linked to antibodies raised againsthuman immunoglobulin. Although widely used for thedetection of antibody against viral (or nonself) antigens,this application was soon followed by studies using themethod to detect autoantibodies. Fluorescent anti-humanglobulin was used in 1957 by Friou and collegues, andHolborow and his co-workers, to demonstrate staining ofthe cell nucleus by serum from patients with systemic lu-pus erythematosus (SLE). These studies, using mouse andhuman tissue sections respectively, also revealed for thefirst time the species nonspecificity of the antinuclear “fac-tor” (ANF), allowing tissue from a wide variety of sourcesto be used. Subsequent studies showed the ANF to be an-tibody, particularly immunoglobulin G (IgG), hence thecurrent terminology of ANA (antinuclear antibody). Fol-lowing closely behind these early studies came reportsthat the pattern of nuclear staining differed between pa-tients with a single autoimmune disease as well as betweenpatients with different autoimmune diseases. Thus homo-geneous staining of the nucleus was more likely in SLE,while nucleolar staining was often found in the serumof patients with scleroderma. During this same period itwas shown that patients with organ-specific autoimmunediseases had serum antibodies which reacted with anti-gens found in the organ targeted by the disease. Thus pa-tients with Hashimoto’s thyroiditis were shown to haveserum antibody directed against antigens in the thyroid.The immunofluorescent test, due to the strong relation-ship between immunofluorescence “pattern” and autoan-tibody specificity, continues to be an important clinicaltest.

2. Principle

The principle of fluorescence is based upon the observa-tion that certain substances adsorb radiation and become“excited” from a ground state of potential energy to thefirst excited state of the molecule. If the excited moleculeis sufficiently stable, it will emit radiation rather than dissi-pate energy to the surrounding molecules as it returns to itsground state. The resulting emission is known as fluores-cence and is almost always of a longer wavelength thanthe exciting radiation. This relationship between excita-tion and emission wavelengths is known as Stokes’ law.

Epifluorescence is the most common method employedin fluorescence microscopy of antibodies. In this techniquethe excitatory radiation is passed through the objectivelens onto the specimen, rather than through the specimen.This means that only reflected excitatory radiation needsto be filtered out to detect the emitted radiation. This is

a significant advantage over transmission of light throughthe specimen, as significantly less irrelevant wavelengthradiation needs to be filtered out. The filtering elementsof an epifluorescence microscope consist of an excitationfilter, an emission filter, and a dichroic mirror (Fig. 2). Theexcitation filter allows transmission of only those wave-lengths that will excite the specific fluorochrome beingused. The emission (or barrier) filter blocks transmissionof the excitatory wavelength light but allows any fluores-cence emitted to pass. The dichroic mirror is coated glassthat is positioned 45◦ to the optical path of the micro-scope. The dichroic mirror functions as a beam splitter,reflecting the excitatory wavelength onto the specimenbut allowing the emitted fluorescence to pass through tothe eyepiece (Fig. 2). In most instances these three el-ements are housed together in a filter cube, with manymicroscopes able to hold two or more filter cubes. Eachset of filters in a cube is a matched set and is restrictedin its use to a small number of fluorochromes, usuallyone.

3. Method

Immunofluorescence is a relatively straightforward tech-nique consisting of four major steps.

a. Preparation of cell or tissue substrates. A va-riety of cell and tissue substrates from numerous specieshave been used in immunofluorescent microscopy to char-acterize both autoantibodies and autoantigens. The mostuseful of these have been transformed mammalian celllines grown in tissue culture. These cell lines contain thegreatest variety of autoantigens and are particularly suitedto the detection of autoantibodies in multisystem autoim-mune diseases where tissue specificity of the autoantibodyis not a confounding consideration. A primary concern inthe preparation of cell or tissue substrates for immunoflu-orescence is that the antigenic integrity of the substrate bepreserved during the fixation procedure necessary to stabi-lize the substrate for experimental use. Another importantaspect of the fixation process is that it permeablizes thecell, thereby allowing antibody to enter and interact withits cognate antigen. Fixation usually involves organic com-pounds, such as ethanol and acetone. Determination of theoptimal fixation conditions for the detection of specificantigens may require considerable experimentation.

b. Addition of primary antibody. Primary antibodymay consist of an unknown specificity, such as that in aclinical sample being tested for the presence of autoan-tibodies, or a known specificity which is being used todetermine the presence of the cognate autoantigen in aparticular cell or tissue substrate.

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FIGURE 2 Function of filter cube elements in epifluorescence microscopy. The three elements of the filter cube, theexcitation filter, emission filter, and dichroic beam splitting mirror, combine to allow selective use of narrow bandwidthsof radiation in conjunction with specific fluorochromes so that fluorescence can be detected. The excitation filter allowsonly certain wavelengths to pass (thick black line), with the dichroic beam splitter reflecting an even narrower bandwidthtoward the specimen. As described by Stokes’ law this excitatory wavelength causes the fluorochrome conjugatedto antibody to emit radiation of a higher wavelength (thick dashed black line), which passes through the dichroicbeam splitter and is transmitted by the emission filter. This radiation is observed as a fluorescent image depicting thelocation in the specimen of the antigen bound by the fluorescently labeled antibody.

c. Addition of secondary antibody. The secondary(or detecting) antibody (antiserum) is immunoglobulin,usually highly purified, that is specific for the species fromwhich the primary antibody originated. Thus if humanserum is being tested for the presence of ANA, then thedetecting antibody will be anti-human immunoglobulins.To be useful in immunofluorescence the secondary anti-body is conjugated to a fluorochrome, most commonly flu-orescein isothiocyanate (FITC) or rhodamine. However, anumber of other fluorochrome are also used, includingAlexa 488 and cyanine (Cy3) dyes, which produce morestable fluorescence.

d. Interpretation of results. Under optimal experi-mental conditions a serum containing an autoantibody willbind to its cognate antigen in the cell or tissue substrate,and this bound antibody will in turn be recognized by thefluorochrome-labeled secondary antibody. When viewedthrough a fluorescence microscope, the emitted fluores-cence identifies the location of the antigen/autoantibodycomplex in the cell or tissue substrate. Success in inter-preting the immunofluorescent “pattern” relies on manyfactors, including the optical properties of the microscope,the specificity and fluorescent label/protein ratio of the

detecting antibody, and the experience of the observer.Considerable uncertainty in “pattern” recognition can beavoided through the use of control sera containing definedautoantibody specificities, such as those available throughthe Centers for Disease Control, Atlanta, Georgia.


1. History

The first report of the use of ELISA (enzyme-linked im-munosorbent assay) to measure antigen specific antibodywas made by Engvall and Perlman in 1972, althoughthe term ELISA had been used earlier by these work-ers to describe a competitive immunoassay for the quan-titative measurement of antigen in solution. Since thoseearly studies ELISA methodology has rapidly expandedto encompass a variety of techniques for the detection ofantigen-specific and nonspecific antibodies, soluble andinsoluble antigens, and cellular antigens, indeed any sub-stance which can generate a specific antibody response.In autoimmunity the technique has become invaluable asa screening assay to detect autoantibodies to a varietyof autoantigens, including proteins and nucleic acids. In

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1990 Burlingame and Rubin described one of the mostinformative analyses of autoantibody reactivity as deter-mined by ELISA. These investigators used subnucleo-some structures as substrates in ELISA to demonstrate thatcomplexes of DNA and histones (e.g., DNA complexed toa dimer of histone 2A and 2B) are better antigens than theindividual components (e.g., histone 2B). These studiesintroduced an additional dimension to the concept that au-toantibodies recognize conformational antigenic determi-nants by suggesting that the quarternary macromolecularstructure may be the target of an autoantibody response.Molecular cloning of autoantigens and their expressionand purification as recombinant proteins have significantlyincreased the usefulness of ELISA in identifying the speci-ficity of autoantibodies in autoimmunity.

2. Principle

In a typical screening assay to detect an antigen-specific(auto)antibody, the antigen is adsorbed onto a solid sup-port, usually a polystyrene microtiter plate. A sourceof soluble antibody (e.g., serum) is allowed to reactwith antigen, unbound material is washed away, andthen an enzyme-conjugated antiimmunoglobulin is added.Unbound (excess) antibody conjugate is removed bywashing, and the amount of antigen-specific antibody de-termined by the addition of an enzyme substrate and mea-surement of the enzyme-generated signal. The signal canbe in the form of a color change as observed when alkalinephosphatase hydrolyzes p-nitrophenylphosphate to pro-duce the yellow p-nitrophenolate. This color change canbe measured using a spectrophotometer at 400 nm. Othermeans of generating signal include enzyme-generated flu-orescence and luminescence.

3. Method

The majority of ELISA procedures that seek to measurethe presence of autoantibodies employ direct adsorption ofautoantigen onto a solid support such as a microtiter plate.The most important aspect of these assays is the purity ofthe antigen. If impure antigen is used, it may be difficult todefine accurately the antigenic specificity of the autoan-tibody. For example, if the antigenic extract contains thenuclear snRNP particles, then both anti-Sm and anti nRNPautoantibodies will be detected. As these autoantibodiescan define different clinical syndromes, any confusion intheir detection is undesirable. On the other hand, the use ofnucleosomes, rather than purified histone, is preferred fordetection of antichromatin autoantibodies. Recent devel-opments in ELISA include the use of purified recombinantautoantigens. Again, these antigenic preparations must becarefully characterized, particularly those from bacterial

expression systems, as most individuals have antibodiesto bacterial components and contamination of recombi-nant protein with bacterial antigens can lead to significant“false-positive” reactions.

C. Immunoblotting

1. History

The term immunoblotting essentially describes thetransfer of antigenic material from one phase to anotherand the use of immunological reagents (i.e., antibody)to detect the transferred protein. The technique has alsobeen called protein blotting and Western blotting, an at-tempt to distinguish the method from the related tech-niques of Southern and Northern blotting, which allowidentification of DNA and RNA, respectively. The termimmunoblotting is more applicable to the technique, par-ticularly when used to characterize the antigenic speci-ficity of autoantibodies. An early and elegant applicationof the procedure was described by Towbin and collegues in1979. They electrophoretically transferred proteins, sepa-rated by polyacrylamide gel electrophoresis, onto a sheetof nitrocellulose (Fig. 3) and then detected the transferredprotein using specific antibody that had been either radio-labeled or conjugated with a fluorochrome or an enzyme.This method exploited the ability of polyacrylamide gelelectrophoresis to separate a mixture of proteins on thebasis of their molecular weights and allowed that level ofresolution to be transferred to nitrocellulose, where spe-cific antibody could be used to identify individual proteins.This method was of immediate applicability to autoim-munity, as contemporary immunological techniques onlyallowed sera to be identified as having the same antigenicspecificity, and did not allow identification of the specificantigen at the molecular level. Within a matter of a fewyears immunoblotting provided not only the molecularweight of many autoantigens, but also clues to the macro-molecular complexity of multicomponent antigens suchas the snRNP particles which contain the Sm and nRNPantigens. As with immunofluorescence, immunoblottingcontinues to be one of the major techniques used to char-acterize both autoantigens and autoantibodies.

2. Principle

Since the early descriptions of the method, immunoblot-ting has undergone considerable modification and manydifferent techniques are in use today (see the Bibliogra-phy for additional reading). The description that followsis essentially the principle of electrophoretic transfer tonitrocellulose from polyacrylamide gels, the techniquepioneered by Towbin, Staehelin, and Gordon in 1979. The

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FIGURE 3 Diagrammatic representation of apparatus used in electrophoretic transfer of proteins from polyacrylamidegel to nitrocellulose. The transfer assembly consists of a “sandwich” made up of an outer porous fiber pad overlaidwith several sheets of adsorbent paper, the polyacrylamide gel, a sheet of nitrocellulose, and then an additional layerof adsorbent paper sheets and another porous fiber pad. Assembly is done with all the components saturated withbuffer to avoid air bubbles that may hinder protein transfer, particularly when between the acrylamide gel and thenitrocellulose sheet. The sandwich is held together by clamps or other firm support and immersed in a buffer-filledtank. Applying voltage results in the proteins being electrophoretically transferred from the polyacrylamide gel to thenitrocellulose. In the case of SDS-PAGE, the negatively charged proteins migrate toward the anode.

exact mechanism that allows proteins to bind to nitrocel-lulose is largely unknown, although hydrophobic forcesmay contribute. Under conditions of sodium dodecylsulfate–polyacrylamide gel electrophoresis (SDS-PAGE),proteins are denatured with SDS and become negativelycharged. The passing of an electric current through thepolyacrylamide gel forces proteins to migrate toward thepositive anode, with the proteins being resolved accord-ing to their molecular weight by virtue of the porosity ofthe gel. Transfer of the resolved proteins to nitrocelluloseoccurs according to essentially the same principle. Underan electric current the negatively charged proteins migrateout of the polycarylamide gel and onto the nitrocellulose(Fig. 3). An important feature of the technique is the ab-

sence of significant SDS in the transfer buffer, as SDS isknown to reduce protein adsorption to nitrocellulose.

3. Method

An immunoblotting protocol consists of numerous steps,from SDS-PAGE resolution of a mixture of antigens tointerpretation of the results (Fig. 4). For simplicity weconsider here only those steps relating to SDS-PAGE res-olution, electrophoretic transfer, and immunological de-tection of autoantigens. The source of protein for SDS-PAGE analysis should be one that contains the antigen orantigens of interest. One means of achieving this is to use acellular substrate that has been found suitable for detection

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Immunology—Autoimmunity 689

FIGURE 4 Immunoblot of rat liver nuclei using human autoim-mune sera. Proteins in purified rat liver nuclei were resolved bySDS-PAGE, transferred to nitrocellulose, and then probed withhuman sera containing autoantibodies to a variety of different au-toantigens. Anti-Pm-Scl serum blots a prominent band at 100 kDa;anti-DNA topoisomerase I (anti-TOPO1) blots a prominent bandbetween 90 and 100 kDa; anti-SS-B/La reacts with a band at48 kDa; antifibrillarin recognizes a band at 34 kDa; antibodiesto nuclear lamins A and C recognize lamin A (higher molecularweight band) and the alternatively spliced lower molecular weightlamin C; antibodies to lamin B1 react with a 69-kDa band. Nu-merical values to the right of the immunoblots represent proteinmolecular weight markers (kDa).

of the antigen using immunofluorescence. For characteri-zation of human autoantibodies extracts from mammaliantissues (e.g., liver) or cell culture lines have been found tobe suitable. If the antigen is only a minor component of thetotal cellular protein (e.g., the nucleolar protein fibrillarin),it may be necessary to purify the appropriate subcellularcompartment (e.g., either the nucleus or the nucleolus) toenrich for the antigen of interest. Electrophoretic transferof autoantigens to nitrocellulose, or other supports, is nomore technically difficult than that of other proteins, how-ever, immunological detection of transferred protein, us-ing autoantibodies, can be challenging. Unlike antibodiesraised by immunization, which are often very high in titer,autoantibody titers can vary considerably between sera. Ifimmunoblotting is being used to detect the presence ofan (auto)antigen, then a high-titer (auto)antiserum should

be used. If the task is to screen a group of sera for au-toantibodies to a particular autoantigen, then care shouldbe taken to ensure that the level of antigen is sufficientand that appropriate “negative” controls are used so that“false-positive” reactions can be eliminated. Considerableuncertainty in recognition of the appropriate bands for par-ticular autoantigens can be avoided through the use of con-trol sera containing relatively monospecific autoantibodyspecificities, such as those available through the Centersfor Disease Control, Atlanta, Georgia. Considerable atten-tion should also be paid to the selection of the secondaryantibody used to detect autoantibody bound to antigen im-mobilized on nitrocellulose. This reagent should be highlyspecific (e.g., affinity purified) and of a high titer. Visual-ization of the autoantigen/autoantibody complex by sec-ondary antibody can be achieved in a variety of ways. Themost sensitive methods include conjugation of secondaryantibody with enzymes such as alkaline phosphatase andhorseradish peroxidase. The addition of appropriate sub-strate allows enzyme-catalyzed colorimetric, fluorescent,or luminescent reactions that can be readily quantified.

D. Immunoprecipitation

1. History

Immunoprecipitation assays can take a variety of forms.Early assays made use of the “immunoprecipitin reac-tion,” which, by mixing of increasing amounts of solubleantigen and antibody, allowed the formation of antigen–antibody complexes. A precipitate would form at the pointof “equivalence,” or when neither antigen nor antibodywas in excess. This type of assay could also be performedin agar or agarose gels (e.g., Oucherlony precipitin test).However, these precipitin assays were cumbersome andrequired large amounts of antigen and antibody to estab-lish optimal conditions for precipitation. The discoverythat components of the bacterial wall (e.g., protein A ofStaphyloccus aureus) could bind the Fc portion of IgGprovided a convenient means of “precipitating” antigen–antibody complexes. Fixed and killed S. aureus could beused to adsorb IgG from serum and then mixed with anti-gen and the resulting antigen–antibody complexes isolatedby centrifugation. Thus purified the properties of the anti-gen and/or antibody could be further examined withoutcontamination from other component of complex mixturesof either antigen or antibody. Covalent coupling of proteinA to Sepharose beads further improved the technique byremoving contaminating bacterial antigens. Immunopre-cipitation using protein A–Sepharose beads has been par-ticularly useful in the characterization of the macromolec-ular structure of autoantigens. Using lysates from wholecells as antigen, and SDS-PAGE to resolve the precipitated

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antigen, it has been found that autoantigens often com-prise complexes of proteins and nucleic acids, such as thesnRNPs of the spliceosome. Precipitation of such macro-molecular complexes is also the major disadvantage ofimmunoprecipitation as it does not allow identification ofindividual antigenic components. This drawback can beovercome by using individual components in immuno-precipitation assays (Fig. 5) or by subjecting the immuno-precipitated complex to immmunoblotting techniques.

2. Principle

Immunoprecipitation is used in autoimmunity to help de-fine autoantibody specificity, as well as to identify thecomponents of the cognate autoantigen. As autoantibod-ies are predominantly of the IgG class, the most com-monly used reagent for immunoprecipitation is protein Abound to Sepharose beads. Protein A interacts with theFc portion of IgG in a reaction that is pH sensitive. Thestrongest interaction occurs in buffers that are neutral orslightly basic in pH, while acidic pH can be used to eluteimmunoglobulin. Not all subclasses of IgG bind to pro-tein A; human IgG3 binds poorly, as does mouse IgG1.Protein A from S. aureus has five IgG binding sites, andprotein A coupled to Sepharose beads binds at least twoIgG molecules. Once an autoantibody-containing serumhas been allowed to react with protein A–Sepharose, theunbound antibody is washed away and a source of antigenadded to the autoantibody–protein A–Sepharose beads.Subsequent identification of the autoantigen is achievedby virtue of prior labeling of the protein and/or nucleic

FIGURE 5 Immunoprecipitation of the autoantigen fibrillarin us-ing autoantibodies. cDNA encoding mouse fibrillarin was radiola-beled with [35S]methionine by in vitro transcription and translation(TnT mFIB). This protein was then used in a protein A–Sepharosebead immunoprecipitation assay to examine human sera (A–L) forantifibrillarin antibodies. Positive sera are identified by an aster-isk. POS. CONT, immunoprecipitate from an antifibrillarin-positiveserum; NEG. CONT., immunoprecipitate from an antifibrillarin-negative serum.

acid components. This is usually achieved by metaboliclabeling of rapidly dividing cell clutures with a radioac-tive precursor such as [35S]methionine for proteins or 32Pifor nucleic acids. The immunoprecipitated antigen is sub-jected to polyacrylamide gel electrophoresis to resolve thecomponents and autoradiography to visualize the radiola-beled components.

Immunprecipitation using extracts from whole cellsmay not allow identification of individual autoantigens,particularly if the autoantigen is a component of a macro-molecular complex. In this case identification of the anti-genic component can be achieved by using the radiola-beled product from the cDNA of the suspected antigen.An example of this antigen-specific immunoprecipitationassay is shown in Fig. 5.

3. Method

As stated above immunoprecipitation of extracts from ra-diolabeled cells has allowed identification of many au-toantigens as components of complexes of protein andnucleic acid (see Tables I and II). As with imunofluores-cence and immunblotting, prior experimentation shouldbe used to confirm the presence of the autoantigen ofinterest in the cell line serving as a source of autoanti-gens. Demonstration of the macromolecular structure ofautoantigens requires considerable experimentation withdifferent conditions of cell lysis and solubilization of cellextract. Conditions that are too stringent can lead to disrup-tion of the complex, while mild conditions may not allowsufficient solubilization to release the complex from sur-rounding cellular constituents. As autoimmune sera cancontain multiple autoantibody specificities, immunopre-cipitation “patterns” revealed by autoradiography can bequite complex. Control sera containing defined autoan-tibody specificities, such as those available through theCenters for Disease Control, Atlanta, Georgia, shouldbe used to help discern the “pattern” of molecular con-stituents of specific autoantigens.


Following the realization that autoantibody specificitiescan serve as diagnostic aids, considerable effort was, andcontinues to be, expended in developing appropriate testsystems for use in research and clinical laboratories. Themajority of these assays focus on the detection of au-toantibody, although the target may be either a singleantigen (e.g., immunoblot, ELISA, imunoprecipitation)or a complex mixture of antigens (e.g., immunofluores-cence, ELISA, immunoprecipitation). Due to the diversityof autoantibody specificities, particularly in multisystem

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autoimmune diseases such as SLE, immunofluorescence(using whole cells as the antigenic substrate) is thescreening assay of choice. Apart from the use of cell cul-ture lines rather than tissue sections, little change has beenmade to the principle of this assay since the 1950s. Fluo-rescence microscopy has seen several advances, includingconfocal microscopy and digitalization of images (includ-ing deconvolution), which have seen considerable use inbasic research but have yet to see use in clinical screeningof autoantibodies. During the last several decades all ofthe methods described have undergone numerous modifi-cations. Most early modifications sought to improve safetyby replacing the use of radioactive detecting reagents(e.g., 125I-labeled secondary antibody) with other meth-ods of detection such as enzyme-catalyzed technologies.These same modifications also improved the sensitivityand shortened the assay time by using colorimetric, fluo-rescent, or luminescent reactions, which are more readilydetected and quantified than radioactive decay. As yet,high-throughput assays have not been applied to autoan-tibody detection, primarily because of the difficulty inscreening an individual serum for the diverse spectrum ofknown autoantibody specificities. However, the availabil-ity of recombinant autoantigens and the increasing arrayof fluorescent or luminescent chromophores suggest thatit should be possible to design assays capable of detectingand quantifying all autoantibodies in any serum using a

complex mixture of autoantigens that are coupled to dif-ferent chromophores.





Chan, E. K. L., and Pollard, K. M. (1997). Detection of autoantibodies toribonucleoprotein particles by immunoblotting. In “Manual of ClinicalLaboratory Immunology,” 5th ed. (N. R. Rose, E. Conway de Macario,J. D. Folds, H. C. Lane and R. M. Nakamura, eds.), pp. 928–934,American Society for Microbiology Press, Washington, DC.

Coligan, J. E., Kruisbeek, A. M., Margulies, D. H., Shevach, E. M., andStrober, W. (eds.) (2000). “Current Protocols in Immunology,” JohnWiley and Sons, New York.

Krapf, A. R., von Muhlen, C. A., Krapf, F. E., Nakamura, R. M., and Tan,E. M. (1996). “Atlas of Immunofluorescent Autoantibodies,” Urbanand Schwarzenberg, Munich.

Peter, J. B., and Shoenfeld, Y. (eds.). (1996). “Autoantibodies,” Elsevier,Amsterdam.

Spector, D. L., Goldman, R. D., and Leinwand, L. A. (eds.) (1997).“Cells: A Laboratory Manual,” Cold Springs Harbor Laboratory Press,Cold Springs Harbor, NY.

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RibozymesAlessandra PoggiJohn J. RossiBeckman Research Institute of the City of Hope

I. IntroductionII. Group I and Group II IntronsIII. Ribonuclease PIV. Self-Cleaving RNAsV. Ribozymes as Tools and Gene Therapy Agents

VI. Conclusions


Cis The same strand of RNA.Functional genomics Identifying the function of genetic

sequences.Gene therapy Supplying therapeutic genes or genetic el-

ements to treat disease.Genotype The genetic makeup of an organism.Introns Sequences interrupting the coding sequence of a

gene or RNA transcript.Phenotype The result of gene expression.Ribozymes RNA molecules with enzymatic activities.Trans A different strand of RNA.


Ribozymes are RNA molecules capable of acting as en-zymes even in the complete absence of proteins. They havethe catalytic activity of breaking and/or forming covalentbonds with extraordinary specificity, accelerating the rateof those reactions. They were simultaneously discovered

in a group I self-splicing intron of the Tetrahymena pre-rRNA and in the ribonuclease (RNase) P enzyme purifiedfrom Escherichia coli. Today there are many additions tothe ribozyme world, including members of the group I andII introns, the self-cleaving domains of the genomes of vi-roids, virusoids, hepatitis D, and a Neurospora satelliteRNA. It has been proposed that the 23S RNA componentof the ribosome may function as a ribozyme in transla-tion, and the U6 snRNA in the spliceosome. Ribozymesoccur naturally, but can also be artificially engineered andsynthesized to target specific sequences in cis or trans.New biochemical activities are being developed using invitro selection protocols as well. Ribozymes can easilybe manipulated to act on novel substrates. These custom-designed RNAs have great potential as therapeutic agentsand are becoming a powerful tool for molecular biologists.

The discovery of catalytic RNA molecules has revo-lutionized views on the origins of life. RNA molecules,once thought to be primarily passive carriers of geneticinformation, can carry out some functions previously onlythought to be catalyzed by proteins, indicating that RNAcan confer not only a genotype (as in many RNA viruses),


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254 Ribozymes

but also a phenotype. The RNA catalyzed reactions in-clude self-cleavage, or trans-cleavage reactions, ligation,and trans-splicing. These observations have led to specu-lation that RNA might have been an early self-replicatingmolecule in the prebiotic world. Evidence supporting thisnotion comes from the fact that group I introns can exhibitRNA polymerase-like activities under certain conditions.In addition, the catalytic core of group I introns shares ho-mology with several small satellite RNAs associated withplant viruses, which are also homologous to the humanhepatitis delta virus (HDV), suggesting a common andancient origin.


Introns are noncoding sequences that interrupt parts ofgenes. When a pre-mRNA is transcribed from the gene,introns need to be removed to give rise to mature mes-sengers that will become a template for protein synthe-sis. Introns are removed by a process of cleavage-ligationcalled splicing. Generally, splicing requires a multicom-plex of proteins and RNA. When introns were first discov-ered in nuclear genes, it was noted that all of their DNAsequences began with a GT and ended with an AG dinu-cleotide (Chambon’s rule). Certain introns isolated fromribosomal or organellar genes, however, did not followthis simple rule, and DNA sequence comparisons led tothe classification of these introns as either group I or groupII on the basis of phylogenetically conserved sequencehomologies and secondary structures. Some examples ofgroup I and II introns are capable of self-splicing in vitroin the absence of protein. Those catalytic RNAs produce5′-phosphate and 3′-OH termini on the reaction products.

A. Group I Introns

Group I introns are widely distributed in fungal mitochon-dria, chloroplasts, rRNA genes of protists, T-even phages,and the genomes of eubacteria. Group I intron self-splicing(in vitro) in the absence of proteins was first observed forthe intervening sequence (IVS, intron) of the nuclear 26SrRNA gene in Tetrahymena thermophila.

Group I splicing proceeds by two consecutive trans-esterification reactions. These reactions are initiated by anucleophilic attack by the 3′-hydroxyl of a guanosine (ora phosphorylated derivative: GMP, GDP, or GTP) at thephosphodiester bond between the 5′-exon and the intron(5′-splice site). The new 3′-hydroxyl group of the 5′-exonthen initiates a second nucleophilic attack, this time on thephosphodiester bond between the 3′-exon and the intron(the 3′-splice site). This results in ligation of the exons andexcision of the intron.

Despite all the evidence for self-splicing in vitro, it isclear that splicing in vivo requires protein factors. Even theTetrahymena IVS, which at low levels of Mg2+ splicesefficiently in vitro, is splicing at a rate of about 50-foldless than the level estimated for splicing in vivo. Proteinstherefore aid in the folding of these complex RNAs toallow the self-splicing reaction to occur.

Self-splicing is, by definition, an intramolecular event,and the intron is therefore not acting as a true enzyme.However, the catalytic activity found within the conservedcore, with a small deletion, can be dissociated into distinctactive enzyme and substrate molecules. Cleavage at the5′- and 3′-splice sites of group I introns can also occurslowly in the absence of a guanosine cofactor, due to thesensitivity of these sites to base hydrolysis which gener-ates cleaved products consistent with the splicing reaction(3′-OH and 5′-P) but unusual for the hydrolysis reaction.The rate of this type of hydrolysis at the splice sites is muchgreater than expected (10-fold higher), implying that thefolded RNA structure influences the susceptibility of cer-tain phosphodiester bonds to alkaline hydrolysis.

Shortened versions of the Tetrahymena IVS (L-19 IVSand L-21 Scal IVS) have been shown to be true enzymesin vitro, for example, as a restriction endoribonuclease andas a template-dependent polymerase.

B. Group II Introns

Group II introns present a relatively restricted distribu-tion; they have been found in plant and fungal mtDNAsand comprise the majority of the introns in chloroplasts.It has also been demonstrated that some members of thegroup II introns can self-splice in vitro. The unimolecu-lar reaction was shown to be Mg2+ dependent, requiringspermidine, having a temperature optimum of 45◦C, andhaving a pH optimum of between 6.5 and 8.5. They dif-fer from group I introns by the structure of their catalyticcore and the intermediate and end products of splicingwhich involve a lariat structure. Again, the reaction con-sists of two transesterifications. Group II introns splice byway of two successive phosphate transfer reactions. In thefirst step, the 2′-OH group of an intramolecular branchpoint adenosine attacks the phosphodiester bond at the5′-splice site (creating a 2′,5′-bond), producing the free5′-exon and a splicing intermediate, the intron-3′-exon.The second step involves cleavage at the 3′-splice site bythe 3′-OH of the 5′-exon. Simultaneously, the exons areligated and the intron lariat, with a 2′,5′-phosphodiesterbond, is released. Base-pairing interactions between se-quences known as the exon binding site (EBS) and theintron binding site (IBS) hold the splice sites in close prox-imity. This ability of group II introns to specifically bindthe 5′-exon has been exploited to encourage the intron to

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catalyze reactions on exogenous substrates. The ability ofthe group II introns to bind the 5′-exon specifically hasbeen exploited to encourage the IVS to catalyze reactionson exogenous substrates. These introns can be engineeredto insert into target RNAs in trans in a reversal of the splic-ing reaction, thereby making them useful for site-specificgene inactivation or site-specific integration of therapeuticgenes.


Ribonuclease P (RNase P) is a ubiquitous endoribonucle-ase that processes the 5′-end of precursor tRNA molecules,producing a 5′-phosphate and 3′-OH termini on the cleav-age products. RNase P consists of both protein and RNAcomponents, and it was shown that the catalyst wasthe RNA moiety. As with the catalytic introns, a diva-lent cation is required as cofactor. RNase P is uniqueamong naturally occurring ribozymes in that it binds andcleaves free substrate molecules; all other characterizedribozymes act in cis. This natural trans-activity makesRNase P an obvious candidate for development as a ther-apeutic agent. Another feature that distinguishes RNaseP from all other ribozymes is that it does not involveWatson–Crick base-pairing between the catalytic RNAand substrate for substrate recognition. Much effort there-fore has been directed toward elucidating the biochemistryand substrate specificity of the RNase P cleavage reaction.One of the first aspects to be analyzed was the role ofthe protein subunit in the cleavage reaction. Comparisonsof the kinetic aspects of the B. subtilis RNA-dependentcleavage reaction performed under various ionic condi-tions have demonstrated that high ionic strength and ad-dition of the protein subunit have similar effects on thekinetics of cleavage. This may indicate that the proteinsubunit acts to disperse the charge repulsions between theRNase P and precursor tRNA substrate RNAs. Becausemany RNase P RNAs are not functional in the absenceof protein in vitro, another possible role for the proteinis to help the RNA moiety to fold into the proper confor-mation. As with most RNA-processing enzymes, the exactsubstrate requirements in terms of sequence and secondarystructure are not well understood. It has been shown thatmature tRNA can compete for binding, suggesting thatmost of the binding energy comes from mature tRNA.Analysis of pre-tRNA deletion mutants showed that onlythe amino acceptor stem and the T loop and stem, whichform a single coaxially stacked helix, are required forcleavage by RNase P, suggesting that any hairpin struc-ture can be cleaved by RNase P provided that a single-stranded NCCA trinucleotide is present at the 3′-side ofthe hairpin. Forster and Altman asked whether a double-

stranded RNA substrate composed of two separate RNAmolecules was also a suitable cleavage substrate. Mixingtwo short complementary oligoribonucleotides containinga 3′-proximal NCCA sequence resulted in cleavage of thetarget RNA at the predicted site. It seems, then, that anyRNA can be cleaved by endogenous RNase P if an exter-nal guide sequence (EGS) containing a single-strandedNCCA at its 3′-end is provided to hybridize with thechosen target.


One category of intramolecular RNA catalysis is thatwhich produces a 2′,3′-cyclic phosphate and 5′-OH ter-minus on the reaction products. A number of small plantpathogenic RNAs (viroids, satellite RNAs, and virusoids),a transcript from a Neurospora mitochondrial DNA plas-mid, and the animal HDV undergo a self-cleavage reactionin vitro in the absence of protein. The reactions requireneutral pH and Mg2+. It is thought that the self-cleavagereaction is an integral part of their in vivo rolling circlemechanism of replication. These self-cleaving RNAs canbe subdivided into groups depending on the sequence andsecondary structure formed around the cleavage site.

A. Hammerhead Ribozymes

This group of RNAs shares a two-dimensional struc-tural motif known as the hammerhead, which has beenshown to be sufficient to direct site-specific cleavage. Thehammerhead structure consists of three base-paired stemswhich flank the susceptible phosphodiester bond, and twosingle-stranded regions, which are highly conserved in se-quence. Extensive mutagenesis has revealed the importantnucleotides and functional groups for efficient catalysis.The hammerhead cleavage domain has been split into twoor three independent RNAs, and trans-cleavage has beendemonstrated in vitro. Haseloff and Gerlach proposed amodel whereby the hammerhead domain is separated suchthat the substrate RNA contains just the cleavage site, andthe ribozyme contains the other conserved nucleotides ofthe catalytic core. Mutagenesis has revealed that the targetsite can be any NUH sequence where H = A, C, and N isany nucleotide. The sequence of the arms of the ribozymealigns the catalytic core to the target site via complemen-tary base-pairing, Analysis has also allowed determinationof the minimum core sequence required for catalytic ac-tivity. The ability to cleave the RNA and thereby inhibitthe expression of a specific gene selectively has two mainapplications: as a surrogate genetic tool for molecular bi-ology and the inactivation of gene transcripts in vivo, asantiviral agents, for example.

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256 Ribozymes

B. Hairpin Ribozyme

A second small catalytic domain is the hairpin structure,which has four helical domains and five loops. Two he-lices of the hairpin domain form between the substrateand ribozyme, and this allows the design and specificityof binding for trans-acting hairpin ribozymes. The hairpinribozyme has a more complicated substrate requirementthan the hammerhead ribozyme, but generally can cleaveto the 5′-side of any GUC.

C. Hepatitis Delta Virus

HDV genomic and antigenomic RNAs contain a self-cleavage site hypothesized to function during rolling circlereplication of this satellite virus. Like the plant pathogens,the sites in HDV are postulated to have a related secondarystructure, three models of which have been proposed:cloverleaf, pseudoknot, and axehead, none of which issimilar to the catalytic domains previously described.Like the other ribozyme motifs, the HDV ribozymes re-quire a divalent cation, and cleavage results in productswith 2′,3′-cyclic phosphate and 5′-OH termini. Investi-gations of trans-cleavage with the HDV ribozyme havenot advanced to those of the hammerhead or hairpinribozymes.

D. Neurospora Mitochondrial VS RNA

The Neurospora mitochondrial VS RNA, a single-stranded circular RNA of 881 nt, shares some features ofthe self-catalytic RNAs of HDV, group I introns, and someplant viral satellite RNAs. Although VS RNA can be de-picted as having a secondary structure like group I introns,it is missing essential base-pairing regions, the cleavagesite is in a different position, and the termini producedare 2′,3′-cyclic phosphate and 5′-OH. Like the hammer-head ribozymes, the VS RNA requires divalent cationsfor cleavage in vitro. The catalytic core of Neurospora VSRNA has been shown to consist of 154 nt.


A. Ribozyme Delivery

Whatever type of ribozyme is chosen, it must be intro-duced into its target cell. Two general mechanisms existfor introducing catalytic RNA molecules into cells: ex-ogenous delivery of the preformed ribozyme and endoge-nous expression from a transcriptional unit. Preformedribozymes can be delivered into cells using liposomes,electroporation, or microinjection. Efforts have been made

to overcome the lack of stability of introduced RNAs byusing modified nucleotides, 2′-fluoro- and 2′-amino-, or2′-O-allyl- and 2′-O-methyl-, mixed DNA/RNA mole-cules, or by the addition of terminal sequences (such asthe bacteriophage T7 transcriptional terminator) at the Y-end of the RNA to protect against cellular nucleases. En-dogenous expression has been achieved by inserting ri-bozyme sequences into the untranslated regions of genestranscribed by RNA polymerase II (pol II), which havestrong promoters, such as the CMV promoter. Ribozymeshave also been inserted into the anticodon loop of tRNA,transcribed by RNA polymerase III (pol III), or have beeninserted into nonessential loop structures of small nuclearRNAs and a small nucleolar RNA. They have been shownto be functional in both plant and animal cells and in trans-genic plants and animals.

Successful use of ribozymes to knockdown target geneexpression is dependent on a number of factors, includingtarget site selection as well as ribozyme gene delivery,expression, stability, and intracellular localization. Thetwo types of ribozymes that have been used most exten-sively for knockdown studies are the hairpin and hammer-head ribozymes. Not all target sites for these ribozymesare accessible for cleavage: secondary structures, bind-ing of proteins and nucleic acids, and other factors influ-ence intracellular ribozyme efficacy. Computer-assistedRNA folding predictions and in vitro cleavage analysesare not necessarily predictive of intracellular or vivo ac-tivity, and the best ribozyme target sites often must bedetermined empirically in vivo. Alternative strategies uti-lizing cell extracts with native mRNAs have also provenuseful for determining accessible ribozyme bindingsites.

Ribozymes can be introduced into cells as genes bytransfection or viral vector transduction, or as chemicallysynthesized molecules stabilized with various base sub-stitutions and 3′ and 5′ modifications. Intracellular deliv-ery, in this case, is typically achieved by some form ofcarrier molecule, such as cationic lipids. Depending onthe application, either delivery method has its advantagesand disadvantages. Delivery of the ribozyme genes canprovide stable intracellular expression and the promoterchoice can allow cell- or tissue-specific expression. Deliv-ery of synthetic ribozymes allows short-term, high-levelavailability, making it easier to use as a “drug;” however,ribozyme stability and pharmacokinetics may present sig-nificant challenges.

Stable intracellular expression of transcriptionally ac-tive ribozymes can be achieved by viral vector-mediateddelivery. Currently, retroviral vectors are the most com-monly used in cell culture, primary cells, and in transgenicanimals. Retroviral vectors have the advantage of stableintegration into a dividing host cell genome, and the

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absence of any viral gene expression reduces the chance ofan immune response in animals. In addition, retrovirusescan be easily pseudo-typed with a variety of envelope pro-teins to broaden or restrict host cell tropism, thus addingan additional level of cellular targeting for ribozyme genedelivery. Adenoviral vectors can be produced at hightiters and provide very efficient transduction, but theydo not integrate into the host genome and, consequently,expression of the transgenes is only transient in activelydividing cells. Other viral delivery systems are activelybeing pursued, such as the adeno-associated virus, alpha-viruses, and lentiviruses. Adeno-associated virus is attrac-tive as a small, nonpathogenic virus that can stably in-tegrate into the host genome. An alphavirus system,using recombinant Semliki Forest virus, provides hightransduction efficiencies of mammalian cells along withcytoplasmic ribozyme expression.

B. Ribozyme Gene Expression

A number of viral promoters (e.g., cytomegalovirus,retroviruses) utilizing RNA pol II have yielded highintracellular expression levels of ribozymes. Furthermore,selectable markers such as antibiotic resistance or cellsurface proteins can be coexpressed with the ribozymeto allow monitoring of ribozyme expression. Embeddingribozymes in a long mRNA transcript can potentiallyhave a negative impact on ribozyme localization andeven folding. One possible solution for this problem isto engineer self-cleaving ribozymes on either side of thefunctional ribozyme so that it can be released from thenascent transcript.

RNA pol III promoters, including tRNA, adenovirusVA1, U1, and U6 small nuclear RNA derivatives, haveallowed ubiquitous and very high intracellular levels ofribozyme expression. Several creative permutations, suchas the combination of two promoters driving the same ri-bozyme gene or the placement of the ribozyme expres-sion cassette within the U3 region of a Long terminalrepeat, to produce a “double-copy” retroviral vectorled to further increases in ribozyme expression. Fi-nally, multiple ribozymes can be expressed within thesame RNA transcript or from multiple promoter-ribozymecassettes.

Intracellular localization of the ribozyme transcriptis another important parameter. Specific strategies tocolocalize ribozymes with their target RNA have beendeveloped to maximize intracellular ribozyme activ-ity. These strategies take advantage of the RNA se-quences flanking the ribozyme. Certain RNA sequencesdirect specific subcellular localization, making it possi-ble to tightly colocalize ribozyme transcripts with theirtarget RNAs.

As the use of ribozymes progresses from cell culturesystems to animal models, additional control over ri-bozyme expression will be required. Ribozyme expressioncan be restricted to specific organs or cell types throughthe use of tissue-specific promoters. This has been donesuccessfully in tissue culture using the tyrosinase pro-moter, which is exclusively expressed in melanocytes. Inanother example, transgenic mice were created that car-ried a ribozyme gene driven by the insulin promoter, onlyexpressed in the pancreatic beta-cell islets. Alternatively,inducible promoters, such as those regulated by tetracy-cline, or steroid hormones, have shown utility both in cellculture and in animals, allowing ribozyme expression tobe turned on and off at will.

C. Ribozyme Applications

Ribozymes have been applied as antiviral agents, treat-ments for cancer and genetic disorders, and as tools forpathway elucidation and target validation. Ribozymes areunique in that they can inactivate specific gene expression,and thus can be used to identify the function of a protein orthe role of a gene in a functional cascade. This application,called target validation, is critical for both basic biologicalresearch and drug development. Compared to other meansof target validation (e.g., transgenic or knockout animals),ribozymes offer specificity and ease of design and usage.

Initial uses of ribozymes focused on antivirals, primar-ily for the treatment of HIV. Viruses that go through a ge-nomic RNA intermediate in their replication cycle, such asHIV, hepatitis B virus, and hepatitis C virus, are attractivetargets because a single species of ribozyme can targetboth viral genomic RNA and mRNAs. Ribozymes havealso been widely used to target cellular genes, includingthose aberrantly expressed in cancers. One early ribozymetarget was the bcr–abl fusion transcript created from thePhiladelphia chromosome associated with chronic myel-ogenous leukemia. This chromosome is characterized by atranslocation that results in the expression of a transform-ing bcr–abl fusion protein. In this case, ribozymes havebeen designed to specifically target the fusion mRNA andnot the normal bcr or abl mRNAs. Ribozymes have alsobeen designed to specifically target the mutant ras genewhile sparing the normal homolog. Ribozymes target-ing overexpressed HER-2/neu in breast carcinoma cellseffectively reduced the tumorigenicity of these cells inmice.

In addition to directly targeting oncogenes, ribozymeshave also been applied more indirectly as anticancer ther-apies. For example, ribozymes targeting the multiple drugresistance-1 or fos mRNAs in cancer cell lines effectivelymade the cells more sensitive to chemotherapeutic ag-ents. Alternatively, a ribozyme targeting bcl-2 triggered

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258 Ribozymes


P1 P2 P3P4



P7P8 P9
















FIGURE 1 The group I intron. The top of this figure depicts the generalized secondary structure of the group I intron.This intron catalyzes the self-splicing of transcripts and can trans-splice as well, as depicted in the bottom portionof the figure. The P1 region forms a helix with the target RNA via Watson–Crick base-pairing, allowing the attachedexon C to be trans-spliced with A.

apoptosis in these cells. Factors required for metastasis arealso attractive targets for ribozymes. Ribozymes targetedagainst CAPL/mts, matrix metalloproteinase-9, pleio-trophin, and VLA-6 integrin all reduced the metastaticpotential of the respective tumor cells in mice. Angio-genesis is also an important target for cancer therapy,and has been blocked in mice by ribozymes targeting fi-broblast growth factor binding protein and pleiotrophin.Ribozyme-based therapies have also been tested in ani-mals to inhibit other proliferative disorders, such as coro-nary artery restenosis.

Heritable and spontaneous genetic disorders representadditional applications for therapeutic ribozymes target-ing cellular genes. These include the beta-amyloid peptideprecursor mRNA involved in Alzheimer’s disease, andan autosomal-dominant point mutation in the rhodopsin

mRNA that gives rise to photoreceptor degeneration andretinitis pigmentosa.

D. Functional Genomics and Target Validation

Ribozymes can be used to inactivate specific gene expres-sion, and thereby can be used to help identify the functionof a protein or the role of a gene in a functional biochemi-cal pathway. Target validation is an increasingly importanttool in basic biological research as well as in drug devel-opment. With the recent completion of the human genomesequencing initiative, there are tens of thousands of tran-scriptomes that have no assigned function. Ribozymesprovide a facile and highly specific tool for interferingwith the expression of these transcripts to monitor theirbiological function.

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5’NNNNNUHNNNNN3’5’NNNNNUHNNNNN3’ | | | | | | | | | | | | | | | | | | | | | |3’NNNNNA3’NNNNNA NNNNN5’ NNNNN5’ AA


YY- - P- - PN--NN--NN--NN--NN--NN--N






Cleavage site

A. Generalized hammerhead ribozyme

FIGURE 2 Generalized hammerhead ribozyme. The ribozyme catalytic core is flanked by stems I, II and III. Cleavageis directed after the H (A, C or U). The N’s represent any nucleotide, Y = pyrimidine, and P = purine. Other bases areA = adenosine, C = cytosine, G = guanosine, and U = uracil.

Ribozyme mediated target validation can also be usedto identify specific member(s) of a protein family involvedin a specific phenotype. Carefully designed ribozymes canselectively knockdown expression of each protein in agene family.

E. Transgenic Animals Expressing Ribozymes

Therapeutics and target validation studies will certainlybe tested in animals. Ribozymes have been used in trans-genic mice to create disease models such as diabetes byselectively downregulating the hexokinase mRNA in pan-creatic islets. In this case, the ribozyme expression wasunder the control of the insulin promoter, and was there-fore only expressed in the pancreatic beta cells. Retroviraldelivery of ribozymes targeted against neuregulin-1 in achick blastoderm resulted in the same embryonic lethal

phenotype as a gene knockout. Localized retroviral deliv-ery of the same ribozyme later in development alloweddissection of the neuregulin biochemical pathway.

F. Ribozyme Mediated RNA Repair

A novel therapeutic application of ribozymes exploits thetrans-splicing activity of the Tetrahymena ribozyme. Thisribozyme has been used to repair defective mRNAs bytrans-splicing onto these RNAs a functional sequence.These ribozymes are designed to bind and cleave the targetRNAs 5′ of the undesired mutation. Since the ribozyme inthis case is an intron, it is engineered to carry with it thecorrect RNA sequence as the 3′-exon. Following cleavageof the mutant target RNA, the ribozyme catalyzes ligationof wild-type sequence onto the cleaved transcript. Thiswas successfully demonstrated with the correction of a

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260 Ribozymes






















H1H1 H2H2



Cleavage siteCleavage site



| | | | | | | | | | | | | | | | | | | |

Generalized Hairpin Ribozyme

FIGURE 3 Generalized hairpin ribozyme. The target RNA (upper sequence) base pairs with the ribozyme (lowersequence) by Watson–Crick base-pairing. Cleavage occurs at the site indicated. The helices formed by the ribozymeand substrate are designated as H1–H4, as indicated. N = any nucleotide, Y = pyrimidine, P = purine. Other basesare A = adenosine, C = cytosine, G = guanosine, and U = uracil.

mutant lacZ transcript, first in bacteria and subsequentlyin correction of a sickle cell message in erythroid cells.

G. Ribozyme Based Gene Discovery

The necessity of ribozymes to selectively base-pair withthe target RNAs prior to catalysis has been exploited todiscover new gene functions involved in a particular phe-notype. This protocol has been termed “inverse genomics.”A combinatorial ribozyme library has been introduced intocultured cells, and selection or scoring for a particular phe-notype has led to the identification of several new genefunctions. A hairpin ribozyme library containing roughly2 × 107 different ribozymes was stably transduced intotumor cells in culture and cells able to grow in soft agarwere identified. Ribozymes isolated from these cells were

sequenced, and the complement to the ribozyme pairingarms was determined. The identity of the target revealeda novel tumor suppressor. This approach has potentialapplications in many systems (therapeutic or otherwise)where a phenotype is of interest but the genes involved areunknown.

H. Ribozyme Evolution

The discovery of the ribozyme sparked new debate on the“RNA world” hypothesis, where all biological processeswere carried out by RNA-based enzymes. Since then,RNA evolution has been forced in vitro to come up withRNA enzymes capable of carrying out a wide variety ofbiochemical reactions, as far-reaching as carbon carbonbond and peptide bond formation. In vitro RNA evolution

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has been used to create RNA-cleaving Rzs with smallercatalytic domains, DNA-cleaving ribozymes, and new cat-alytic motifs. Even RNA-cleaving DNAzymes have beengenerated through in vitro evolution. These “evolved”enzymes exemplify the power of in vitro evolution andwill no doubt find many applications.


The transformation of ribozyme sequences from naturallyoccurring, cis-cleaving molecules to target-specific, trans-cleaving reagents has stimulated a great deal of interestin their potential applications. Ribozymes targeting viralgenes are now in clinical evaluation; ribozymes targetingcellular genes are moving into transgenic animals; andthe use of ribozymes is expanding into RNA evolution,mRNA repair, and gene discovery.

For ribozymes to become generally useful surrogategenetic tools and realistic therapeutic agents, several ob-stacles need first to be overcome. These obstacles are theefficient delivery to a high percentage of the cell popula-tion, efficient expression of the ribozyme from a vectoror intracellular ribozyme concentration, colocalization ofthe ribozyme with the target, specificity of ribozyme forthe desired mRNA, and an enhancement of ribozyme-mediated substrate turnover. Despite these reservations,results with ribozymes so far look promising, particularlyin the HIV-1 studies. As our knowledge of RNA struc-ture, secondary and tertiary, increases, we will be able totarget RNAs more rationally, which may help with theproblems of specificity. At the same time, the understand-ing of the physical localization of RNA in cells and itstracking as it moves from the nucleus to cytoplasm willalso help in ensuring colocalization of the ribozyme andtarget. Modifications of the ribozymes, for example, the2′-ribose with various agents such as methyl, allyl, flu-oro, and amino groups, increases the stability to nucle-ases quite dramatically. Similarly, chimeric DNA–RNAribozymes increase the stability. The efficiency of deliv-ery to cells with viral vectors or liposomes is also contin-

ually improving. These molecules must retain their cat-alytic potential, reach an accessible site on the substrate,and effectively impact on the steady-state levels of targetmolecules to be useful as either surrogate genetic tools ortherapeutic agents. Great progress has been made in all ofthese areas and should allow extensive use of the highlyspecific reagents for down-regulating expression of targetRNAs.





Castanotto, D., Rossi, J. J., and Deshler, J. O. (1992). “Biological andFunctional Aspects of Catalytic RNAs,” Crit. Rev. Eukaryot. GeneExpr. 2, 331.

Cech, T. R. (1990). “Self-Splicing of Group I Introns,” Annu. Rev.Biochem. 59, 543.

Doherty, E. A., and Doudna, J. A. (2000). “Ribozyme Structures andMechanisms,” Annu. Rev. Biochem. 69, 597.

Fedor, M. J. (2000). “Structure and Function of the Hairpin Ribozyme,”J. Mol. Biol. 297, 269.

Hauswirth, W. W., and Lewin, A. S. (2000). “Ribozyme Uses in RetinalGene Therapy,” Progr. Retin. Eye Res. 19, 689.

James, H. A. (2000). “Therapeutic Potential of Ribozymes in Haemato-logical Disorders,” Expert Opin. Investig. Drugs 9, 1009.

Kurz, J. C., and Fierke, C. A. (2000). “Ribonuclease P: A Ribonucleo-protein Enzyme,” Curr. Opin. Chem. Biol. 4, 553.

Rossi, J. J. (1999). “Ribozymes, Genomics and Therapeutics,” Chem.Biol. 6, R33.

Rossi, J. J. (2000). “Ribozyme Therapy for HIV Infection,” Adv. Drug.Deliv. Rev. 44, 71.

Szostak, J. W. (1997). “In Vitro Selection and Directed Evolution,” Har-vey Lect. 93, 95.

Watanabe, T., and Sullenger, B. A. (2000). “RNA Repair: A Novel Ap-proach to Gene Therapy,” Adv. Drug Deliv. Rev. 44, 109.

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Translation of RNA to ProteinR. A. CoxH. R. V. ArnsteinNational Institute for Medical Research

I. IntroductionII. mRNA Structure and the Genetic CodeIII. Transfer RNAIV. Ribosome Structure and Function in TranslationV. Translational Control of Gene Expression

VI. Concluding Remarks


Anticodon Three consecutive bases in tRNA that bind toa specific mRNA codon by complementary antiparallelbase-pairing.

Antiparallel base-pairing Pairing through specific hy-drogen bonds between base residues of two polynu-cleotide chains or two segments of a single chain withphosphodiesterbonds running in the 5′ → 3′ directionin one chain or segment and in the 3′ → 5′ directionin the other. In DNA and RNA the hydrogen bondsare usually formed between complementary base pairs(A with either T or U, and G with C).

Elongation The stepwise addition of amino acids to thecarboxyl terminus of a growing polypeptide chain.

Initiation A multistep reaction between ribosomal sub-units, charged initiator transfer RNA, and messengerRNA that results in apposition of the ribosome-boundinitiator Met-tRNA with an AUG initiator codon inmRNA. In this position, the ribosome is poised to formthe first peptide bond.

Polarity The asymmetry of a polymer such as a polynu-

cleotide or polypeptide. In DNA, the two strands haveopposite polarity; that is, they run in opposite direc-tions (5′ → 3′ and 3′ → 5′). The polarity of a polypep-tide is defined as running from the N-terminus to theC-terminus.

Reading frame One of three possible ways of trans-lating groups of three nucleotides in mRNA. The ap-propriate reading frame is determined by the initiationcodon.

Template strand The strand of the DNA double helixthat is used as a template for transcription of RNA.It has a base sequence complementary to the RNAtranscript.

Termination The end of polypeptide synthesis, which issignaled by a codon for which there is no correspondingaminoacyl-tRNA. When the ribosome reaches a termi-nation codon in the mRNA, the polypeptide is releasedand the ribosome-mRNA-tRNA complex dissociates.

Translation The stepwise synthesis of a polypeptidewith an amino acid sequence determined by the nu-cleotide sequence of the mRNA coding region. Thegenetic code relates each amino acid to a group of three


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32 Translation of RNA to Protein

consecutive nucleotides termed a codon. Decoding ofmRNA takes place in the 5′ → 3′ direction, and thepolypeptide is synthesized from the amino to the car-boxyl terminus.

Translocation The stepwise advance of a ribosome alongmRNA, one codon at a time, with simultaneous transferof peptidyl-tRNA from the A site to the P site of theribosome.

PROTEINS are essential to the structure and functionof living cells. The assembly of polypeptide chains fromamino acids and their subsequent modifications, leadingto the final three-dimensional protein structure, are ex-ceptionally complex processes; many components are in-volved and much of the cell’s energy is utilized. Eachpeptide bond requires the expenditure of four high-energyphosphate bonds. This value excludes the energy usedfor initiation and release of the polypeptide chains andthe cost of synthesizing and processing mRNA. The lin-ear amino acid sequence of a protein is encoded withinthe gene as a linear deoxyribonucleotide sequence. Earlysteps in the biosynthesis of a protein include transcrip-tion of the gene and appropriate processing of the tran-script leading to the production of mature messengerRNA (mRNA). We describe the mechanisms involvedin translating mRNA to produce a polypeptide chainwhich has the amino acid sequence specified by thegene.


A gene or cistron is defined as the region of DNA thatis transcribed into a functional RNA. The transcript func-tions either as such (e.g., tRNA, rRNA, snRNA) or as amessenger (mRNA), which, after processing or editingas required, normally codes for one or more polypep-tide chains in the translation process. A polynucleotidesuch as RNA is an asymmetrical polymer assembledfrom nucleoside triphosphates by a stepwise mechanismlinking the 5′ position of one nucleotide by a phos-phate bridge to the 3′ position of the adjacent nucleotide.In the finished polynucleotide chain the first nucleotideresidue has a 5′ position which is not linked to an-other nucleotide, whereas the last nucleotide has an un-linked 3′ position. Thus, polynucleotide synthesis pro-ceeds from the 5′ to the 3′ terminus and the polymeris said to have a 5′ to 3′ polarity. Usually, linear RNAsequences are written with the 5′ terminus on the leftand the 3′ terminus on the right (Fig. 1A). Within theRNA chain some bases may form antiparallel base pairs(Fig. 1B).

The polypeptide chains of proteins are also asymmetri-cal polymers in which the amino acid residues are linkedby peptide bonds between their alpha amino and carboxylgroups (Fig. 2), leaving a free alpha amino group at oneend (the amino terminus) of the polymer and a free alphacarboxyl group at the opposite end (the carboxyl termi-nus). The significance of the polarity of RNA and proteinswill become evident when the process of protein biosyn-thesis is explained below.

The genetic information stored in DNA is not usabledirectly for making proteins. Rather, it must be copiedinto a primary RNA transcript containing the mRNA by anenzymatic transcription of segments of DNA containingthe genes. In prokaryotes, the primary transcript is also themessenger; that is, it can be used directly in polypeptidesynthesis. In contrast, in eukaryotes the primary transcriptis often much larger in size than the mature mRNA andrequires extensive processing involving the excision ofintervening and other noncoding sequences.

Messenger RNA serves as the template for protein syn-thesis; that is, the linear nucleotide sequence of the mRNAdictates the amino acid sequence of the polypeptideencoded originally by the gene. Conventionally, geneand mRNA nucleotide sequences are written in the 5′

to 3′ direction, which corresponds to the direction inwhich mRNA is decoded during polypeptide synthesis:The mRNA is read in the 5′ to 3′ direction, and thepolypeptide is synthesized from the amino- toward thecarboxyl-terminus.

The mechanism whereby RNA is translated into proteinis complex, and the cell devotes considerable resources tothe translational machinery. The components include 20different amino acids, transfer RNAs, aminoacyl-tRNAsynthetases, ribosomes, and a number of protein factorswhich cycle on and off the ribosomes and facilitate varioussteps in initiation of translation, elongation of the nascentpolypeptide chain, and termination of synthesis with re-lease of the completed polypeptide from the ribosome.The process depends on a supply of energy provided byATP and GTP. The rate of protein synthesis is typicallyin the range of 6 (immature red blood cells of the rabbit)to 20 (Escherichia coli growing optimally) peptide bondsper sec. at 37◦C.


A. Structure

The sequence information of a gene is copied (transcribed)into the nucleotide sequence of RNA from the com-plementary strand of DNA, called the template strand.The primary transcript is a single strand of RNA, which

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FIGURE 1 Structural elements of ribonucleic acids. (A) Primary structure indicating the numbering system for purines,pyrimidines, and ribose. (B) Base-pairing interactions commonly found in RNA: (i) G + C base pair; (ii) A + U basepair; (iii) G + U base pair. Pairs i and ii are Watson and Crick base pairs; iii is a special type of base pairing found inintramolecular bihelical regions. The minor groove is on the side of the base pair with the glycosidic bond of whichthe carbon atom C1′ of ribose is boxed. Note that DNA contains base pairs of types i and ii only but with thymine(5-methyluracil) in place of uracil. [From Arnstein, H. R. V., and Cox, R. A. (1992). “Protein Biosynthesis,” OxfordUniversity Press, Oxford. With permission.]

is a faithful copy of the other strand of DNA with sub-stitution of U residues in place of T residues found inDNA. Sometimes, the primary transcript is altered, as de-scribed below, before it functions as mRNA; in these cases,the original unmodified transcript is the precursor or pre-

mRNA. Usually, mRNAs have nontranslated sequences atthe 5′ and 3′ ends in addition to the coding domain. Thesenoncoding sequences sometimes affect the efficiency oftranslation and the stability of mRNA. The structures oftypical mRNAs are shown in Fig. 3. The decoding process

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34 Translation of RNA to Protein

FIGURE 2 Structure of the peptide bond. (a) Two L-amino acidswith different side chains, R1 and R2; (b) A dipeptide formed fromthe two amino acids shown in (a). [From Cox, R. A., and Arnstein,H. R. V. (1995). In “Encyclopedia of Molecular Biology and Mole-cular Medicine” (R. A. Meyers, ed.), Volume 6, pp. 108–125. VCHPublishers, New York. With permission.]

involves base-pairing between three bases (designated acodon) in the mRNA and the three-base anticodon of atransfer RNA (tRNA). In a separate reaction, each tRNAis first linked to a particular amino acid and thus the pairingof mRNA with tRNA determines the sequence of aminoacids in the resulting protein.

FIGURE 3 Structure of typical mRNAs. (a) Prokaryotic mRNA. At the beginning of the transcript there are two ad-ditional phosphate residues linked by pyrophosphate bonds to the 5′ phosphate group of the terminal nucleotide(see Fig. 1). The 5′ untranslated sequence often contains a ribosome binding site (the Shine–Dalgarno sequence)which increases the efficiency of translation. AUG and UAA are representative initiation and termination codons,respectively. Cistrons (gene sequences) are separated by short (typically 12 to 24 nucleotides) noncoding inter-cistronic sequences. (b) Eukaryotic mRNA. Typically, the processed transcripts are monocistronic. The 5′ end isusually modified by the addition of 7-MeG, known as a cap structure, which is linked by two pyrophosphate groupsto the terminal nucleotide of mRNA. The coding sequence is located between 5′ and 3′ untranslated sequences.In the majority of cases the 3′ end is modified by the addition of 25 to 250 adenylate residues, termed the poly(A)tail. [From Arnstein, H. R. V., and Cox, R. A. (1992). “Protein Biosynthesis,” Oxford University Press, Oxford. Withpermission.]

B. Prokaryotic mRNA

In organisms that do not have a nucleus (prokaryotes), pre-mRNA usually undergoes little or no modification so thatpre-mRNA and mRNA are very similar if not identical. Be-cause pre-mRNA is colinear with DNA, DNA and proteinsare usually colinear in these organisms. Gene expressionin prokaryotes usually involves the co-transcription of sev-eral adjacent genes, and translation of mRNA sequencesinto polypeptides may begin at the 5′ end of mRNA whiletranscription is still in progress at the 3′ end.

C. Eukaryotic mRNA

In cells with a nucleus (eukaryotes) the genetic informa-tion is stored mainly in the nucleus and to a minor degreein some organelles (mitochondria and chloroplasts). Thedescription that follows pertains only to nuclear genes.Eukaryotic genes are more complicated than prokaryoticgenes because the coding region is often discontinuous:the coding sequences or exons are interrupted by inter-vening sequences (introns). Thus, genes and proteins areusually not colinear in eukaryotes. In the nucleus, a com-plicated set of splicing reactions removes all the intronsfrom pre-mRNA and fuses the exons into a continuouscoding sequence. Other processing steps involve adding a“cap” to the 5′ end of the mRNA and a poly(A) “tail” tothe 3′ end. After completion of these nuclear maturationsteps the mRNA is transported to the cytoplasm where it is

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translated. As with prokaryotic mRNA the coding regionis flanked by 5′ and 3′ nontranslated sequences.

D. The Genetic Code

The genetic code is triplet, comma-less, and nonoverlap-ping. As a consequence, a nucleotide sequence has threepossible reading frames (Fig. 4). Because mRNA is nor-mally translated into a unique polypeptide, an essentialstep in the translation process is the selection of the ap-propriate reading frame. This is achieved by starting trans-lation at the initiation codon, usually AUG or less fre-quently GUG, which ensures that the following codonsare read in phase within the required reading frame. Ofthe 64 theoretically possible triplets in the genetic code,61 sense codons correspond to the 20 genetically encodedamino acids found in all, or nearly all, proteins. WhenGUG is used as the initiation codon, it codes for me-thionine by interacting with the anticodon of the initiatorMet-tRNAMet, whereas elsewhere it codes for valine. Allother codons specify only one amino acid but many aminoacids are specified by two or more (up to six) codons; thecode is unambiguous, but degenerate. The remaining threecodons, termed nonsense codons, usually signify termina-tion of synthesis and release of the finished polypeptidechain.

1. Deviations from the Standard Genetic Code

One of the nonsense codons, UGA, has an additionalfunction in the synthesis of selenoproteins. The processinvolves the initial synthesis of selenocysteyl-tRNA froma novel seryl-tRNA and selenium. This tRNA contains ananticodon that is able to decode UGA and insert seleno-cysteine residues into the growing polypeptide chain, but

FIGURE 4 Translation of a polynucleotide sequence into three al-ternative polypeptides using different reading frames. [From Cox,R. A., and Arnstein, H. R. V. (1995). In “Encyclopedia of MolecularBiology and Molecular Medicine” (R. A. Meyers, ed.), Volume 6,pp. 108–125. VCH Publishers, New York. With permission.]

only at UGA codons in a particular context of neighboringnucleotides. The insertion of selenocysteine residues intothe polypeptide also requires a specific elongation factorT which differs from the factor used for the incorporationof other aminoacyl-tRNAs.

Mitochondria and chloroplasts, as well as certain organ-isms such as mycoplasma and ciliated protozoa, use a fewnonstandard genetic codons. For example, methionine isusually coded for by AUG (or occasionally by GUG) butin human mitochondria this codon is replaced by AUA.Variations in the genetic code are thought to have arisenas a result of the loss of some tRNA genes and mutationalpressure on DNA, giving rise to a predominance of eitherAT- or GC-rich codons.


By relating individual codons of mRNA to the cognateamino acids, tRNA functions as a key bilingual interme-diate in the translation of the genetic code. All transferRNAs are single-stranded molecules about 80 nucleotideslong with a common 3′-terminal CCA sequence. Most ofthe bases are standard but some (e.g., pseudoU, dihydroU,and T) are derived by modification after transcription ofthe transfer RNA genes.

The secondary structure of tRNA is usually presentedin two dimensions as a cloverleaf to highlight the regionsof base-pairing (Fig. 5a). X-ray crystallography revealsthat additional hydrogen bonds give rise to an L-shapedtertiary structure (Fig. 5b). The CCA sequence carryingthe amino acid is located distal to the anticodon.

The decoding process involves antiparallel base pairingbetween the three bases of mRNA codons and the comple-mentary anticodons of transfer RNA (tRNA) during pep-tide bond formation (Fig. 6). The first and middle bases

∗Transfer RNA nomenclature: The amino acid linked to a chargedtRNA is indicated by a prefix and the specificity of the transfer RNA(tRNA) in the aminoacylation reaction is shown as a superscript on theright; for example, Phe-tRNAPhe indicates phenylalanine-specific tRNAcharged with phenylalanine. The anticodon may be indicated as a rightsubscript or, alternatively, in the superscript after the amino acid; forexample, tRNAUGC or tRNAAla/UGC. The right-hand subscript positionis sometimes used to indicate the organism from which the tRNA isderived (e.g., tRNAVal

yeast). The initiator tRNA, which is specific for me-thionine, is termed tRNAMet

f or tRNAMeti . In the cytosol of eukaryotes the

charged initiator tRNA is termed Met-tRNAMetf of Met-tRNAMet

i . Oftenthe superscript Met is omitted. In prokaryotes and in the mitochondriaof eukaryotes, the methionine residue of the charged initiator tRNA isformylated by a transformylase using N10-formyltetrahydrofolate as thedonor, giving N-formylmet-tRNAf. Commonly, this charged tRNA istermed fMet-tRNAf. The methionine-specific elongator RNA, which in-serts methionine into internal positions of the growing peptide chain,is termed tRNAMet

m (or tRNAm) when uncharged, and Met-tRNAMetm

(or Met-tRNAm) when charged.

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FIGURE 5 Structure of phenylalanyl-transfer RNA. (a) Sec-ondary cloverleaf structure. The 5′ and 3′ ends of the moleculeare marked; the continuous line represents the sugar phosphatebackbone. The short lines denote base residues and the dots de-note base-pairing through standard hydrogen bonds (see Fig. 1).The D loop contains dihydrouracil residues; the T�C loop containsthymine and pseudouridine. (b) Tertiary structure. The abbrevia-tions for unusual bases are defined in (a). The sugar phosphatebackbone is represented by double parallel lines. Standard basepairs are represented by short double lines, and nonstandardbase pairs by single lines. The anticodon sequence is stippled,and the acceptor end is shaded. [From Arnstein, H. R. V., and Cox,R. A. (1992). “Protein Biosynthesis,” Oxford University Press, Ox-ford. With permission.]

FIGURE 6 Schematic illustration of base pairing between acodon and its anticodon. The diagram shows the interaction at theP site of the ribosome (see Fig. 9) between the initiation codonAUG of mRNA and the anticodon CAU of fMet-tRNAMet

f and the in-teraction at the A site between the codon GUA of mRNA and theanticodon UAC of Val-tRNAVal. The polarity of an RNA speciesruns from the 5′ end to the 3′ end. The fragment of tRNA is rep-resentative of the general structure (see Fig. 5b) placed in theappropriate orientation; the numbers refer to the nucleotide po-sitions measured from the 5′ end. The interaction between thecodon and the anticodon is antiparallel, and the three base pairshave a bihelical conformation.

of the codon form conventional base pairs with the thirdand middle bases of the anticodon, respectively, but thethird base of the codon pairs with the first base of the anti-codon by a less stringent interaction (e.g., base-pairingof G with U as well as with C), giving rise to degen-eracy. This so-called “wobble” considerably reduces thenumber of tRNA species required to decode the 61 sensecodons. Thus, the protein synthesis system in the cytosolof eukaryotes contains only a few more than 40 differenttRNAs, and in mitochondria 22 to 24 tRNA species aresufficient.

The attachment of amino acids to tRNA involves the for-mation of an ester bond between the alpha-carboxyl groupof the amino acid and the 3′-hydroxyl group of the termi-nal adenosine of tRNA. It requires specific enzymes, theaminoacyl-tRNA synthetases. There are 20 different syn-thetases, each specific for one of the 20 amino acids, andeach enzyme recognizes something unique in the structureof its cognate tRNA. The structural determinants whichensure accuracy of this charging reaction vary for differenttRNAs. The anticodon may play a part but sometimes evena single base elsewhere is sufficient to determine the speci-ficity of the tRNA-synthetase interaction. The accuracy of

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Translation of RNA to Protein 37

the synthetase reaction in attaching an amino acid to itscognate tRNA is critically important to the fidelity of thetranslation process. Once the aminoacyl-tRNA has beenformed, the subsequent incorporation of the amino acidresidue into a polypeptide does not depend on the aminoacid itself but only on the interaction between the anti-codon of the aminoacyl-tRNA with the codon of mRNA.Thus, an error in the synthetase reaction would lead tothe incorporation of an inappropriate amino acid into thepolypeptide.

Synthesis of aminoacyl-tRNA (III) from amino acids(I) requires activation of the amino acid carboxyl groupwith formation of an intermediate enzyme-bound amino-acyladenylate (II).

E + R−



−CO−2 +ATP → E · R−



−CO · AMP + PPi


(I) (II)

→ R −



−CO · tRNA + AMP + E


where E = aminoacyl-tRNA synthetase.The energy for the reaction (two high-energy phosphate

bonds) is provided by ATP and stored in the ester bond ofthe aminoacyl-tRNA to be used subsequently for peptidebond synthesis.


A. Ribosome Structure

Ribosomes are high-molecular-weight complexes of RNA(rRNA) and proteins (Table I), and the electron-dense par-ticles are easily visualized by electron microscopy. Ri-bosomes from various sources (prokaryotes, eukaryoticcytoplasm, mitochondria, chloroplasts, and kinetoplasts)vary in size from 20 to 30 nm in diameter, but all are com-posed of a large and a small subparticle or subunit andperform similar functions in protein synthesis. The prin-cipal functional domains of the ribosome and associatedcomponents are given in Fig. 10. More detailed resolutionof the ribosome structure has allowed the placement ofmRNA, aminoacyl-tRNA, peptidyl-tRNA, and the nascentpolypeptide chain (see Section C and Fig. 11).

The small subunit comprises a single rRNA of 0.3–0.7 × 106 Da and single copies of 20 to 30 unique proteins.It has a major function in binding initiator tRNA andmRNA in the initiation of protein synthesis and in de-coding the genetic message.

The large subunit comprises a high-molecular-weightrRNA (0.6–1.7 × 106 Da) and often one or two smallerrRNAs (0.03–0.05 × 106 Da) and 30 to 50 different pro-teins are present, with one exception, as single copies.The large subunit binds aminoacyl-tRNA at the A-site,peptidyl-tRNA at the P site, and discharged tRNA at theE (exit) site. The large subunit contains the peptidyl trans-ferase and, unusually, this enzyme activity resides in therRNA molecule itself rather than in the associated ribo-somal proteins. This subunit is also involved in bindingelongation factor G which is required for translocation(Fig. 10).

B. The Ribosome Cycle in Translation

Polypeptide synthesis can be divided into three stages:initiation, elongation, and termination (see Fig. 7). Initia-tion involves the binding of a ribosome to mRNA with theinitiation codon correctly placed in the P-site. Elongationleads to the stepwise increase in the length of the polypep-tide chain through the transfer of the growing chain to theamino group of aminoacyl-tRNA. Termination of chainelongation and release of the completed polypeptide oc-curs when a termination codon reaches the A site. Allstages require the participation of protein factors. Ad-vances in establishing the structures of translational fac-tors by X-ray crystallography, which have been rapidin the last decade, are reviewed by Al-Karadaghi et al.(2000).

1. Formation of Pre-Initiation Complexes

The ribosome cycle starts with the stepwise formationof an initiation complex from mRNA, charged initiatortRNA, and ribosomal subunits. A number of pre-initiationcomplexes are formed as intermediates and the processis facilitated by initiation factors. In outline, prokaryoticand eukaryotic systems are similar, but there are a fewdifferences, particularly as regards the complexity of theinitiation factors and details of the mechanisms.

a. Prokaryotic systems. Three proteins, initiationfactors IF-1, IF-2, and IF-3 (see Table II), are requiredfor the initiation of protein biosynthesis (see Fig. 8a).Ribosomal subunits are released by dissociation of ribo-somes following translation of the mRNA. Dissociation isfacilitated by the combined action of the initiation factorsIF-1 and IF-3; IF-1 increases the rate of dissociation and

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38 Translation of RNA to Protein

TABLE I Properties of Ribosomes and Ribosomal Subunits

RNA:protein rRNA S20,w

Source S20,w Size (nm)a Mass (Mda) (w/w) Axial ratio mass (Mda) (nominal)


E coli 70 22.5 ± 2.5 2.6–2.9 2:1 — f


Cytoplasm 80 28.0 ± 2.8 3.4–4.5 1:1 —

Chloroplast 70 22.5 ± 2.5 2.5–3.3 1:1 —


Protistsc 60–80 ca. 3.25 1:1.38 —

Animals 55–60 2.7–3.2 1:1.7–1:4 —

Fungi 67–80 4.2 1:1.3–1:1.6 —

Higher plants 78 — — —

Small ribosomal subunits


E. coli 30 22.0 ± 2.2 0.95 2:1 2:1 0.6 16


Cytoplasm 36–41 25.0 ± 2.5 1.4 1:1 2:1 0.7 18

Chloroplast 28–35 22.0 ± 2.2 1.2 1:1 2:1 0.6 16


Protists 32–45 — — — 0.2–0.35 11–16

Animals 32–40 1.1–1.6 — — 0.31 12–14

Fungi 32–29 1.6–1.7 — — 0.45–0.64 15–19

Higher plants 44 — — — 0.64 18

Large ribosomal subunits


E. coli 50 22.5 ± 2.2 1.75 2:1 1:1 1.17 23

0.04 5


Cytoplasm 60 28.0 ± 2.8 2.1–3.1 1:1 1:1 1.2–1.75 25–28

0.05 5.8

0.4 5

Chloroplasts 46–54 22.5 ± 2.2 2.4 1:1 1:1 1.1 23

0.04 5

0.03d 4.55d


Protists 45–60 — — — 0.4–0.74 12–24

Animals 25–35 1.65–2.0 — — 0.5 16–21

Fungi 50 — — — 0.77–1.22 21–25

Higher plants 60 — — — 1.25 26

0.04e 5e

a Largest dimension.b Mitochondrial ribosomes are preferentially associated with the inner mitochondrial membrane and are more diverse than

cytoplasmic or chloroplast ribosomes. Particular features include post-transcriptional oligoadenylation of the 3′ ends of both thesmaller and larger rRNA components. Except for mitochondrial ribosomes of higher plants, the essential sequence motifs 5SrRNA and 5.8S rRNA are present in the large rRNA component and are not found as individual species.

c Mitochondria of protists such as Trypanosoma brucei are also known as kinetoplasts.d 4.5S rRNA corresponding to the 100 nucleotides at the 3′ end of the 23S rRNA in eubacteria occurs as a separate species in

chloroplasts of higher plants.e This species is present only in plant mitochondria and is absent from all other mitochondrial ribosomes.f Data not available.

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TABLE II Prokaryotic Initiation Factors from E. coli


Factor (kDa) Properties and function

IF-1 9 Stimulates activity of IF-2; accelerates dissociationof unprogrammed ribosomes to subunits.

IF-2 100 Binds fMet–tRNAf to the ribosomal P site by aGTP-requiring reaction.

IF-3 22 Binds natural mRNAs to the small ribosomalsubunit probably by facilitating base-pairingbetween the untranslated leader sequence andthe 3′ end of 16S rRNA; prevents ribosomalsubunit association when bound to thesmall subunit.

From Arnstein, H. R. V., and Cox, R. A. (1992). “Protein Biosynthe-sis,” Oxford University Press, London. With permission.

IF-3 acts as an anti-association factor when bound to the30S ribosomal subunit, thereby displacing the equilibriumin favor of subunit formation. Initiation factor IF-2 is alsoable to bind to the 30S subunit and this association isstabilized by IF-1 and GTP, the latter acting as a stericeffector without being hydrolyzed at this stage. IF-2plays a central role in binding fMet-tRNAf to the 30Spre-initiation complex by specific recognition of the N-formylmethionine residue attached to the initiator tRNA,thus restricting this interaction to charged initiator tRNA.All three factors bind to the 30S ribosomal subunit nearthe 3′ end of the 16S ribosomal RNA at adjacent sites thatare located at the interface between the small and largeribosomal subunits.

In the next step, the initiator tRNA and mRNA as-sociate with the 30S–IF-1–IF-2–IF-3 complex with re-lease of IF-3. There is evidence from in vitro experimentsthat the binding of mRNA precedes that of the initiatortRNA.

Messenger RNA binds to the small ribosomal subunitimmediately before formation of the final initiation com-plex with the initiation codon correctly positioned in theP-site (see Fig. 7a). In the case of bacterial and bacte-riophage messengers, the molecular recognition mecha-nism proposed by Shine and Dalgarno (1974) involvesbasepairing between short nucleotide sequences, most of-ten CUCC, near the 3′ end of the 16S ribosomal RNAand a complementary region, usually consisting of 3 to 9bases on the 5′ side of the mRNA initiation codon, whichhas been found to be present in nearly all of more than150 bacterial and bacteriophage messengers. Studies withmutants and mRNA fragments indicate that, in additionto the Shine–Dalgarno interaction, outlying upstream se-quences in the leader region may also provide recognitionsignals between mRNAs and ribosomes, possibly by en-suring that the Shine–Dalgarno sequence is in an appro-priate conformation.

The Shine–Dalgarno mechanism is also found inchloroplast protein synthesis as judged from sequenceanalysis of the 16S rRNA and mRNAs, but apparentlynot in the mammalian mitochondrial system where theinitiator codon occurs either directly at, or only a few nu-cleotides downstream from, the 5′ end of mRNA, whichexcludes the possibility of mRNA–rRNA base-pairing inthis region.

b. Eukaryotic systems. At least 12 proteins, the eu-karyotic initiation factors (eIF) (see Table III), are neededfor initiation of protein biosynthesis (Fig. 8b). The dis-sociation of cytosolic 80S ribosomes is facilitated by acomplex initiation factor, eIF-3 (Mr approx. 5–700,000),consisting of 9 to 11 polypeptide chains, which binds tothe small ribosomal subunit (40S) and prevents its re-association to 80S ribosomes. Thus, this factor has anti-association activity, but low-molecular-weight proteinswith similar activity have also been reported, and a protein,eIF-4C, of Mr 20,000, seems to function as an accessoryfactor to eIF-3 in the formation of a 43S ribosomal pre-initiation complex. Also, another protein factor, eIF-6, ofMr 24,000, prevents re-association by binding to the large(60S) ribosomal subunit.

Initiation factor eIF-2 gives a stable binary com-plex with GTP which binds the initiator tRNA, Met-tRNAf, forming a ternary complex. Interaction of thisternary complex with the 40S ribosomal subunit contain-ing bound initiation factors eIF-3 and eIF-4C gives riseto the 43S pre-initiation complex, which is competentto bind messenger RNA in the presence of three furtherinitiation factors, eIF-4A, eIF-4B, and eIF-4F, togetherwith ATP.

The binding of cytosolic eukaryotic messenger RNAsto the small ribosomal subunit probably does not involvebase-pairing with the 18S rRNA, as no uninterrupted se-quences of the Shine–Dalgarno type have been found.Instead, a “scanning model” has been proposed, in whichthe pre-initiation complex, composed of the 40S ribosomalsubunit, Met-tRNAMet

f and associated initiation factors,binds at or near the 5′ cap of the mRNA and slides alongthe messenger until it encounters the first AUG triplet,at which point the 60S ribosomal subunit joins to giverise to the 80S initiation complex. Recognition of the capis facilitated by cap-binding proteins, which mediate anATP-dependent melting of the mRNA secondary struc-ture at the 5′-terminal region to allow the mRNA to threadthrough a channel in the neck of the 40S subunit. Thecap structure is required for efficient binding and transla-tion even in cases where the initiating AUG codon occurshundreds of nucleotides downstream. As a rule, scanningby the 40S subunit stalls at the first AUG codon, whichis recognized mainly by interaction with the anticodon

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40 Translation of RNA to Protein

FIGURE 7 Schematic view of the biosynthesis of a polypeptide by translation of prokaryotic mRNA. (a) Formationof the initiation complex from mRNA, ribosomal subunits, initiation factors (IF), GTP, and fMet-tRNAMet (for details,see Fig. 8). The binding sites for aminoacyl-tRNA, peptidyl-tRNA, and uncharged tRNA are designated A, P, and E,respectively. (b) Decoding of the codon GUA with Val-tRNAVal. (c) Formation of the peptide bond by transfer of fMet toVal-tRNAVal forming fMet–Val-tRNAVal. (d) Translocation of tRNAMet

f to the E site and of fMet–Val-tRNAVal to the P sitewith codon UUC aligned with the A site. (e) Ejection of tRNAMet

f from the ribosome. (f) Decoding of codon UUC withPhe-tRNAPhe. (g) Decoding of codon UAA with release factor. Steps (b) to (f) constitute the elongation–translocationcycle. The position shown is reached by repeating the cycle n + 2 times after formation of fMet. Val-tRNA. (h) Release ofthe completed polypeptide, fMet–Val–Phe–(aa)n–Ser, ribosomal subunits, release factor, mRNA, and tRNASer. Thecycle may then be repeated starting at (a). [From Cox, R. A., and Arnstein, H. R. V. (1997). In “Encyclopedia ofMolecular Biology and Molecular Medicine,” (R. A. Meyers, ed.), Volume 6, pp. 108–125. VCH Publishers, New York.With permission.]

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FIGURE 8 Schematic diagram illustrating the formation of initiation complexes. (a) Prokaryotic initiation. The followingsymbols are used for initiation factors and other components involved in the formation of the 70S initiation complex:�, IF-1; �, IF-2; �, IF-3; , fMet-tRNAf, �, GTP; other symbols are defined in the figure. (b) Eukaryotic initiationcomplexes in mammalian protein biosynthesis from components defined in the figure. The process may be dividedinto three stages: a, formation of a 43S pre-initiation complex; b, binding of mRNA with formation of a 48S pre-initiationcomplex; and c, synthesis of the 80S initiation complex containing the initiator tRNA in the correct position for peptidebond formation. [From Arnstein, H. R. V., and Cox, R. A. (1992). “Protein Biosynthesis,” Oxford University Press,Oxford. With permission.]

of the Met-tRNAMetf . However, this recognition also de-

pends in some way on eIF-2 and may be modulated as aresult of deviation of the mRNA structure from the consen-sus sequence GCCGCCA/GCCAUGG. Sequence contextmay also account for the rare cases where initiation occursdownstream of the 5′-proximal AUG codon. Where this

sequence context is unfavorable, initiation becomes ineffi-cient, hence most 40S subunits will tend to initiate furtheralong the mRNA at another AUG triplet in a more favor-able context. This model also explains rare cases whereinitiation is not restricted to one particular AUG codon andtranslation of a single mRNA gives rise to two proteins.

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TABLE III Eukaryotic Initiation Factors

Initiation Mr (kDa) of factorsfactor Synonym and subunits Properties and function

eIF-1 15 Stabilizes initiation complexes

eIF-2 alpha, 38a GTP-dependent binding of Met-tRNAf to the small ribosomal subunitbeta, 35–50a

gamma, 55

eIF-2B GEF 27, 37, 52, 67, 85a Conversion of eIF-2–GDP into eIF-2–GTP

eIF-3 9–11 subunits Associates with 40S subunit to maintain dissociation; binds mRNA to24–170a 43S preinitiation complex

eIF-4A 50-kDa component 50 ATP-dependent unwinding of the secondary structure of the mRNA 5′of CBP-II region; stimulates translation of exogenous mRNA in cell-free systems

eIF-4B 80a mRNA binding; stimulates cell-free translation; ATPase activity of eIF-4Aand eIF-4F; AUG recognition and recycling of eIF-4F

eIF-4C 17 Ribosome dissociation; 60S subunit joining

eIF-4D 17 Formation of first peptide bond

eIF-4E CBP-I; 24-kDa CBP 24–28a Binds mRNA cap structure

eIF-4F CBP-II; cap-binding 24 (CBP-I),a ATPase; unwinds mRNA secondary structure; stimulates cell-freeprotein complex 50 (eIF-4A), 200a translation

eIF-4G 220 Stimulates protein synthesis by interacting with eIF-4E and poly(A)protein to circularize polysomes

eIF-5 60a GTPase; release of eIF-2 and eIF-3 from pre-initiation complex to allowjoining of the 60S subunit

eIF-6 24 Anti-association activity; binds to the 60S ribosomal subunit

a Denotes subunit can be phosphorylated in vivo.Abbreviations: GEF, guanine nucleotide exchange factor; CBP, cap-binding protein.

2. Initiation Complex Formation: Joiningof the Large Ribosomal Subunit

The last event in the initiation of protein synthesis in-volves the joining of the large ribosomal subunit to thepre-initiation complex (Fig. 8). In the prokaryotic sys-tem, association of the 50S subunit with the 30S pre-initiation complex takes place with hydrolysis of GTP bythe GTPase activity of IF-2 and release of IF-1, IF-2, GDP,and Pi. GTP hydrolysis is essential for the release of IF-2from the initiation complex, which is a prerequisite for al-lowing the fMet-tRNAf to engage in the formation of thefirst peptide bond. In eukaryotic protein synthesis, the 80Sinitiation complex is formed by joining the 60S ribosomalsubunit to the 48S pre-initiation complex consisting of the40S ribosomal subunit, eIF-2, eIF-3, GTP, Met-tRNAf,mRNA, and possibly eIF-4C. This coupling reaction re-quires an additional factor, eIF-5, which mediates the hy-drolysis of GTP to GDP with release of eIF-2-GDP, Pi,and eIF-3 from the 48S pre-initiation complex.

By this stage, all initiation factors have been releasedand are available for recycling, although the exact steps atwhich factors are released from intermediate complexesare not known in every case. There is thus an initiationfactor cycle within the ribosome cycle, and regulation ofthe activity of factors, particularly eIF-2, is an importantcontrol mechanism in translation (see Section V.D.1).

In the initiation complex, location of the charged initia-tor tRNA in the P site of the ribosome allows transferof the methionine residue to the amino group of an-other aminoacyl-tRNA in the A site (Fig. 7b) by pep-tidyl transferase to form dipeptidyl-tRNA (see Fig. 7c).Functional insertion of Met-tRNAf directly into the P sitecan be demonstrated using the trinucleotide AUG as asynthetic mRNA and another trinucleotide, for exampleUUU, to bind an acceptor aminoacyl-tRNA (in this casePhe-tRNA).

It is possible to measure the peptidyl transferase activityof the large subunit in the absence of mRNA by usingthe antibiotic puromycin, which resembles the 3′-terminalregion of Phe-tRNA in structure, as an artificial acceptorto form methionyl puromycin from Met-tRNAf.

3. Polypeptide Chain Synthesis: The Elongation–Translocation Cycle

This cycle is outlined in Figs. 7b–f.

a. Elongation. The first peptide bond is formed whenthe aminoacyl-tRNA in the ribosomal A site is convertedinto the corresponding methionyl-aminoacyl-tRNA bytransfer of the methionyl (or N-formylmethionyl) residuefrom the charged initiator tRNA in the P site (Fig. 7c). In

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artificial cell-free systems, any N-substituted aminoacyl-tRNA, such as peptidyl-tRNA or N-acetylaminoacyl-tRNA, can function in peptide bond synthesis as a donorin the P site in place of the charged initiator tRNA. Thereaction is catalyzed by the peptidyltransferase activity ofthe large ribosomal subunit. No soluble cofactors appearto be involved, but monovalent cations (K+) at a concen-tration of 100 mM or more and divalent cations (Mg2+)below 2 mM are required.

Efficient entry of aminoacyl-tRNA into the ribosomalA site requires the participation of an elongation factor,termed EF-Tu in prokaryotes (see Table IV) and EF-1(EF-1L) in eukaryotes (see Table V), and GTP. This elon-gation factor forms a ternary complex with GTP and allaminoacyl-tRNAs except initiator tRNA, but not withuncharged tRNA, thus ensuring that only appropriatelycharged tRNAs are efficiently bound in the A site. A spe-cial elongation factor showing extensive homology withboth EF-Tu and IF-2 is involved in the synthesis of seleno-proteins (see Section II.C) from selenocysteyl-tRNAUCA

in E. coli.The above-mentioned aminoacyl-tRNA binding reac-

tion catalyzed by EF-Tu is the rate-limiting step in theelongation cycle; peptide bond formation and transloca-tion are much faster. The initial binding of the ternarycomplex to the ribosome is readily reversed, but the inter-action is stabilized by the subsequent codon recognitionwhich induces the GTPase conformation of EF-Tu lead-ing immediately to the hydrolysis of the GTP componentof the ternary complex to GDP. Hydrolysis of the GTPmoiety causes a further change in the conformation of EF-Tu from the GTP-binding to the GDP-binding form. Thisconformational change leads to the release of aminoacyl-tRNA, allowing its CCA end to align with the peptidyltransferase center of the ribosome and the instantaneous

TABLE IV Properties of Prokaryotic Elongation and Termi-nation Factors from E. coli

Mr (kDa) Properties and function

Elongation factors

EF-Tu 43 N-terminal acetyl-serine; heat labile; bindsaminaocyl-tRNA to the ribosomal A site

EF-Ts 30 Heat stable; regeneration of EFTu–GTP

EF-G 77 GTP-dependent translocation of peptidyl-tRNAand its mRNA codon from the A site to theP site of the ribosome

Termination (release) factors

RF1 36 Requires UAA or UAG codons for hydrolysisof peptidyl-tRNA

RF2 38 Requires UAA or UGA codons for hydrolysisof peptidyl-tRNA

RF3 46 Enhances RF1 and RF2 activity

TABLE V Eukaryotic Elongation and Termination Factors

Mr (kDa) Properties and function

Elongation factors from various yeast, animal, and plant cells

eEF-1A (EF-1L 50–60 Analogous to EF-Tu

or eEF-Tu)

eEF-1B (eEF-Ts) 30 Analogous to EF-Ts

eEF-2 105 Contains essential SH groups andone residue of a post-translationallymodified histidine residue;GTP-dependent translocationanalogous to EF-G

eEF-3 125 GTPase and ATPase activity; functionnot fully defined

Termination or release factors

eRF-1 110 Two 55-kDa subunits; binds to theribosome A site by a GTP andtermination codon dependentreaction; hydrolyzes peptidyl-tRNAin the P site

eRF-2 GTPase; stimulates eRF-1 activity

formation of the peptide bond. The elongation factor islater released from the ribosome as a complex with GDP.The operation of EF-Tu is thus similar to that of initiationfactor 2 which binds charged initiator tRNA to the smallribosomal subunit.

Following dissociation from the ribosome the EF-Tu–GDP complex interacts with another elongation factor,EF-Ts, with formation of an EF-Tu–EF-Ts heterodimerand release of GDP. Reaction of the heterodimer withGTP regenerates the EF-Tu–GTP complex required forbinding aminoacyl-tRNA. The sequence of events is simi-lar in eukaryotes with eEF-1A (Mr 50,000) correspondingto EF-Tu and eEF-1B (Mr 30,000) to EF-Ts.

Selection of the specific aminoacyl-tRNA to be boundat the ribosomal A site is by base-pairing between therelevant mRNA codon and the tRNA anticodon. Becausethis interaction involves only a triplet of bases and hencea maximum of nine hydrogen bonds (see Fig. 1B), it isintrinsically unstable at physiological temperatures andis probably stabilized by components of the ribosome toallow sufficient time for peptide bond synthesis to occur.Also, the codon–anticodon pairing must be monitored forfidelity in order to minimize errors in translation. In E. colithere is genetic and biochemical evidence that one of theproteins of the small ribosomal subunit, S12, is involved inensuring the fidelity of normal translation and in causingthe mistranslation which occurs in the presence of theantibiotic streptomycin due to incorrect codon-anticodoninteractions.

b. Translocation. Translocation involves the move-ment of the ribosome along the mRNA in the 5′ → 3′

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direction. Immediately after synthesis of the first pep-tide bond, the ribosomal A site contains dipeptidyl-tRNAwhile uncharged initiator tRNA remains in the P site.Thus, both these sites are occupied, and to allow the nextaminoacyl-tRNA to enter the A site it is necessary to ejectthe uncharged tRNA and shift the dipeptidyl-tRNA fromthe A into the P site. This translocation (Fig. 7d) takes placeas a concerted process involving movement of both mes-senger RNA and dipeptidyl RNA together into the P site,leaving the A site occupied by the next mRNA codon andfree to accept the cognate aminoacyl-tRNA (see Fig. 7e).At the same time, the deacylated tRNA moves first intoan E (exit) site with subsequent ejection when the nextaminoacyl-tRNA enters the A site.

Translocation requires the participation of another elon-gation factor (EF-G in prokaryotes and EF-2 in eukary-otes) and GTP (Table IV). It seems that when EF-G andGTP bind to the ribosome, translocation occurs but GTPhydrolysis is required only subsequently to release EF-Gand GDP. The location of the EF-G binding site on the ri-bosome overlaps with that for EF-Tu, thus EF-G must bereleased before the EF-Tu–aminoacyl-tRNA–GTP com-plex can enter the A site. Analogous reactions occur ineukaryotic systems.

There is little information about the details of the trans-location mechanism. A continuous polyribonucleotidechain is not essential, as translocation can occur with in-dividual trinucleotides. It seems likely that movement ofthe mRNA is dependent on and tightly coupled to thatof the tRNA with the binding sites for the tRNA provid-ing the precision for movement by exactly one codon.Presumably, binding of EF-G and GTP after release ofEF-Tu–GDP following peptide bond synthesis induces aconformational change in the ribosome which leads totranslocation.

After translocation the ribosomal P site is occupiedby dipeptidyl-tRNA and the vacant A site contains thethird mRNA codon. Entry of the next aminoacyl-tRNA,selected as before by the codon–anticodon interaction,into the A site (Fig. 7f) enables peptide bond synthe-sis to continue and repeated operation of the elongation-translocation cycle gives rise to a stepwise elongation ofthe nascent polypeptide chain, each complete cycle elon-gating the chain by one amino acid residue and movingthe mRNA by one codon in the 5′ to 3′ direction. When theend of the coding sequence is reached and one of the termi-nation (or stop) codons has entered the A site, translationstops and the completed polypeptide chain is released.

c. Termination. (See Figs. 7g–h.) The presence ofone of the three termination codons, UAA, UAG, or UGA,in the A site results in the binding of a release factor(Table IV) instead of an aminoacyl-tRNA to the ribosome.

In prokaryotes, two release factors have been identified,one (RF1) recognizing UAA and UAG, the other (RF2)functioning with UGA. Ribosomal binding and release ofRF1 and RF2 are stimulated by a third factor, RF3, whichinteracts with GTP and GDP. In eukaryotic cells such asreticulocytes, one release factor (eRF) has been found tofunction with all three termination codons, and the bindingof this factor to ribosomes is stimulated by GTP but notGDP. Although the details are not entirely clear, GTPhydrolysis appears to be required for the release of thefinished polypeptide chain by cleavage of the peptidyl-tRNA bond and completion of the termination processleading to dissociation of the release factor from theribosome.

Thus, at the end of the ribosome cycle the coding se-quence of messenger RNA has been translated to producea particular polypeptide chain, and all the components in-volved become available for re-use in another round ofthe cycle (Fig. 7h). Usually, several ribosomes becomeattached to one mRNA molecule, giving rise to polyribo-somes (also called polysomes; Fig. 9). In eukaryotic cellsthe efficiency of protein synthesis is stimulated by factoreIF4-G (Table III), which interacts with both factor eIF-4Eand a poly A-binding protein. The resultant circularizedpolysomes show an enhanced ability to re-initiate after re-lease of the ribosomal subunits from the messenger RNAat the end of a round of translation.

C. High-Resolution Structural Studiesof the Ribosome

Electron microscopy of ribosomes has provided sufficientinformation to allow the construction of models showingthe general features of the principal functional domains,such as the location of mRNA, ribosomal subunits, andfactors (Fig. 10).

During the last ten years, X-ray diffraction studies haveled to considerable advances in the elucidation of the struc-ture of the ribosome in more detail, culminating in the de-termination of the E. coli ribosome at 0.78-nm resolution(Fig. 11) and of the small and large subunits at resolu-tions of 0.3 nm and 0.25 nm, respectively. The structureshave revealed the identity of each amino acid and each nu-cleotide. The findings provide insights at the atomic levelinto the reactions leading to the decoding of mRNA andthe formation of the peptide bond. Moreover, the impor-tance of the role of rRNA in ribosome function has be-come evident from the structures; the functional regionsof both small and large subunits are rich in RNA. Thethree-dimensional structures also highlight the dynamicaspects of ribosome function leading to the view that theribosome is a highly sophisticated motor driven by GTPwith rRNA playing a leading role.

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FIGURE 9 Electron micrograph of a thin section through twoneighboring epithelial cells showing endoplasmic reticulum (largearrows) to which are attached numerous polysomes. Groups offree ribosomes (small arrows) occur in the cytoplasm; pm, plasmamembrane. Bar represents 1000 nm. [Micrograph by I. D. J.Burdett, National Institute for Medical Research, London, U.K.]

The higher resolution studies of subunits have also re-vealed the mode of action of several antibiotics such asparomomycin, streptomycin, and spectinomycin, whichmodify the decoding function of the small subunit, andchloramphenicol, puromycin, and vernamycin, which af-fect peptide bond formation.


Cells needs to synthesize specific proteins in the requiredamounts at particular times and to deliver them to the cor-rect locations. These processes depend on a great manyinteractions and numerous mechanisms exist for the trans-lational control of gene expression. Examples of the waysused by cells to control the translation of mRNA are pre-sented in the sections that follow. This material is intendedto be illustrative rather than comprehensive, and it is to beexpected that novel control mechanisms will continue tobe discovered.

FIGURE 10 Model of the E. coli ribosome, based on low-resolution electron microscopy. The diagram shows the relativeorientation of the large and small subunits and other functionalcomponents involved in polypeptide chain synthesis. H, head ofthe small subunit; CP, central protuberance and L7/L12 stalk ofthe large subunit; EF-Tu, elongation factor required for bindingaminoacyl-tRNA to the ribosomal A site; EF-G, elongation factorrequired for translocation of the peptidyl-tRNA from the A to the Psite. The broken line indicates the boundary between the transla-tional and exit domains which are involved in peptide bond forma-tion and extrusion of the nascent polypeptide chain, respectively.[From Cox, R. A., and Arnstein, H. R. V. (1997). In “Encyclope-dia of Molecular Biology and Molecular Medicine” (R. A. Meyers,ed.), Volume 6, pp. 108–125. VCH Publishers, New York. Withpermission.]

A. mRNA Stability

Provided all other components of the translational systemsare present in optimum amounts, control of translationmay be achieved through the availability of the relevantmessenger RNAs at the site of protein synthesis. Thesteady-state level of mRNA is determined by the rate ofits synthesis and degradation. Control of transcription isof major importance for synthesis in both prokaryotes andeukaryotes. The stability of mRNA depends on its pri-mary and secondary structure as well as on the presenceof factors such as stabilizing proteins and nucleases.

The secondary structure of mRNA is determined by itsnucleotide sequence. For any mRNA a number of codingsequences are possible because of the degeneracy of thegenetic code. Thus, degeneracy allows for particular fea-tures of secondary structure (often termed cis factors) to

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FIGURE 11 Electron density of the 70S ribosome. (A) Stereo view of the 0.78-nm electron density map from thesolvent side of the 30S subunit (purple). The 30S subunit consists of the head (H) connected to the platform (P) andbody (B). Additional features of the small subunit are the neck (N), spur (SP), shoulder (S), and contacts betweenthe head and platform (a and b). The 50S subunit (gray) has the prominent features of the protein L1 stalk, centralprotuberance (CP), and L7/L12 region. (B) View of the ribosome rotated 90◦ about the vertical, with the L7/L12 regionin the foreground. The A-site finger is marked with an asterisk. (C) View of the ribosome rotated 180◦ about the verticalaxis compared with (A). (D) Model of three tRNAs bound to the ribosome, viewed toward the 30S interface (left) andtoward the 50S interface (right). All three tRNA surfaces are electron density maps calculated to 0.78-nm resolution.The 3′-CCA end of the A-site tRNA (marked with ∧) is not modeled for lack of electron density to constrain its position.The 3′-CCA end of the E-site tRNA (asterisk) is buried at the base of the L1 stalk and cannot be distinguished at thepresent resolution. A, P, and E denote A, P, and E site, respectively. The peptidyl transferase center is located on thelarge subunit near the lower contact point of the A- and P-site-bound tRNAs. [From Cate, J. et al. (1999). Science285, 2095–2104. With permission.]

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be favored, which may act either to stabilize or destabilizethe mRNA, according to the needs of the cell, by determin-ing its susceptibility to degradative enzymes (often termedtrans-acting factors).

Whereas the structure of mRNA determines its suscep-tibility to degradative enzymes, the detailed mechanismsare complex. In prokaryotes, the enzymes involved includetwo endonucleases (RNase E and RNase III) and two ex-onucleases (polynucleotide phosphorylase and RNase II).Other nucleases may be active in particular cases such asphage infection. In eukaryotes, a major pathway involvesremoval of the 3′ poly(A) tail (deadenylation), followedby removal of the 5′ cap, which renders the mRNA suscep-tible to rapid endonucleolytic degradation in the 5′ → 3′


B. Control by Interaction of Proteinswith mRNA

Throughout the ribosome cycle, dynamic protein–mRNAinteractions are functionally important in the initiation,elongation, and termination of polypeptide synthesis.In addition, more stable associations between proteinsand mRNAs have been observed, particularly in eukary-otic cells. These messenger ribonucleoprotein complexes(mRNPs) occur both in polyribosomes and free in thecytosol, some of the latter being either temporarily orpermanently unavailable for translation. Thus, protein-mRNA interactions contribute to the efficiency with whichmRNAs are translated.

Some proteins, such as the poly(A)-binding protein(p78), are present in most if not all mRNPs, whereas othersappear to be cell specific and mRNA selective. In unfer-tilized sea urchin eggs and Xenopus oocytes, for example,untranslated messenger is sequestered by association withproteins that prevent translation until later stages of devel-opment. Duck reticulocytes contain globin mRNP, whichcannot be translated in vitro, whereas the mRNA obtainedby deproteinizing the complex can be translated, show-ing that in this case translation is prevented by the mRNPproteins.

Formation of a site-specific mRNA-protein complex isinvolved in the translational control of the biosynthesisof ferritin, an iron storage protein, which is stimulated inresponse to the presence of iron. In this instance, a cy-toplasmic repressor protein of 85 kDa binds to a highlyconserved 28-nucleotide stem–loop structure in the 5′ un-translated region of ferritin mRNAs in the absence of iron.In the presence of iron, the protein dissociates from themRNA, which is then available for translation. A similarloop motif occurs in the 3′ untranslated region of transfer-rin receptor mRNA, which is also subject to translationalcontrol by an iron-responsive repressor.

During the cell cycle, histone mRNA is destabilizedafter completion of DNA replication, resulting in a 30-to 50-fold decrease. This change appears to be due to anincrease in the level of free histones, which form a com-plex with histone mRNA by interaction with a stem–loopstructure at the extreme 3′ terminus. Formation of thishistone–histone mRNA complex is thought to activate aribsome-associated 3′ → 5′-exonuclease, which degradesthe histone mRNA. During the S phase, newly synthesizedDNA binds free histones to form nucleosomes, thus pre-venting the degradation of histone mRNA at this stage ofthe cell cycle.

Specific regulation of gene expression at the level oftranslation also exists in prokaryotes. For example, thesynthesis of E. coli threonyl-tRNA synthetase is negativelyautoregulated by an interaction of the tRNA-like leadersequence of its mRNA with the synthetase, which in-hibits translation by preventing the binding of ribosomes.The synthetase is displaced from the mRNA by tRNAThr,which thus acts as a translational antirepressor. This reg-ulatory mechanism allows the cell to maintain a balancebetween the tRNA synthetase and its cognate tRNA.

Similarly, there is a mechanism used to control thesynthesis of proteins encoded by a polycistronic mRNA.In this case, selective binding of the ribosomal proteinto the region of the mRNA involved in the initiation oftranslation leads to the regulatory protein controlling bothits own synthesis and that of other ribosomal proteins.A specific example is the role of ribosomal protein S4,which acts as a translational repressor of four riboso-mal proteins (S4, S11, S13, and L17). Protein S4 appearsto function as a repressor through an unusual “pseudo-knot” linking a hairpin loop upstream of the ribosome-binding site with sequences 2 to 10 codons downstreamof the initiation codon. (A pseudoknot structure containsintramolecular base pairs between base residues in theloop of a stem–loop structure and distal complementaryregions of the RNA.) Stabilization of this structure by S4would prevent the binding of ribosomes, and this controlmechanism may contribute to the coordinated synthesisof the different ribosomal proteins required for ribosomeassembly.

C. Control by mRNA Structure

The secondary structure of some eukaryotic mRNAs reg-ulates translation by a mechanism involving a riboso-mal frameshift which gives rise to a directed changeof the translational reading frame to allow the synthe-sis of a single protein from two or more overlappinggenes by suppression of an intervening termination codon.Several retroviruses use this mechanism to move from onereading frame to another in the expression of the viral

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FIGURE 12 Translational control of bacteriophage synthesis. (a) Arrangements of MS2 and f2 bacteriophage cistrons.(b) Arrangement of Qβ bacteriophage cistrons. (c) Replicative intermediate synthesizing new plus strands on thecomplementary minus strand copy of the original bacteriophage RNA. The ribosome binding sites (RBS) are indicatedby the numbered arrows. The major RBS (1) binds ribosomes efficiently but can be blocked by ribosomal protein S1.The secondary RBS (2) becomes available only after translation of at least part of the coat protein cistron by ribosomes.RBS 3 is masked by the secondary structure of native bacteriophage RNA but is accessible in nascent RNA (c) orin vitro when the secondary structure is destroyed. Noncoding regions are shown in black upstream from the RBSsites and at the 3-end. [From Arnstein, H. R. V., and Cox, R. A. (1992). “Protein Synthesis,” Oxford University Press,Oxford. With permission.]

RNA-dependent DNA polymerase. Other example of theoperation of such a frameshift include the synthesis ofthe reverse transcriptase enzymes of several retrotrans-posons, such as the yeast Ty1. The mechanism of chang-ing the reading frame involves “slippery” sequences anda complex folding of the mRNA into a structure termed apseudoknot.

In E. coli, translational control of the three cistrons ofQβ , f2, and related bacteriophage RNAs (Fig. 12) accountsfor the synthesis of coat protein:replicase:A protein inthe approximate ratio of 20:5:1. These quantitative differ-

ences are due to the differential and independent initiationof translation at each cistron as a result of differences inthe secondary structure of the mRNA initiation sites. Fur-thermore, in vivo there is a delay in the synthesis of coatprotein and this temporal control involves translationalrepression of the cistron by ribosomal protein S1. In ad-dition, S1 functions as one of the subunits of the f2 RNAreplicase; therefore, association of the newly synthesizedtranslation product of the f2 replicase cistron with S1 willfavor its dissociation from the phage RNA, thus allowingtranslation of the coat protein cistron to start.

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D. Control by Modification of TranslationFactor Activity

1. Initiation Factors

In eukaryotes, initiation of protein synthesis is inhibitedby phosphorylation of the initiation factor eIF-2. Phos-phorylation is stimulated by the lack of haem or thepresence of double-stranded RNA. Two different pro-tein kinases capable of phosphorylating the alpha sub-unit of eIF-2 have been characterized. One enzyme, calledthe haem-controlled repressor (HCR) or haem-regulatedinhibitor (HRI), is a cytoplasmic protein (95,000 Da) thatbecomes activated by phosphorylation. The other kinase(67,000 Da) is activated by phosphorylation in the pres-ence of double-stranded RNA. Thus, a cascade of pro-tein phosphorylation is involved. Phosphorylated eIF-2is unable to exchange GDP for GTP, which preventsit from functioning in the binding of initiator tRNA toribosomes.

Conditions other than lack of haem (e.g., heat shock,serum deprivation, or the presence of oxidized glu-tathione), which are known to inhibit protein synthesis,also give rise to the phosphorylation of eIF-2α. Con-versely, the activity of eIF-4F is decreased by dephospho-rylation of the 24-Da subunit (see Table III) rather than byphosphorylation. Thus, a number of different kinases andphosphatases are involved in modulating the activities ofdifferent factors.

Small RNAs may also be involved in regulating thetranslation of mRNA in eukaryotic cells. Of the stimula-tory RNAs, the best characterized is a small RNA of about160 nucleotides, which accumulates in cells after infectionwith adenovirus. This virus-associated RNA, VA-RNA1,which is required to maintain general protein synthesis,acts by inhibiting the phosphorylation of the alpha sub-unit of initiation factor eIF-2.

2. Other Translation Factors

A Ca2+/calmodulin-dependent protein kinase phospho-rylates eEF-2. The phosphorylated factor appears to beinactive and moreover also inhibits the activity of the non-phosphorylated factor. Dephosphorylation of the factor byphosphatase restores its activity.

E. Effects of Antisense Polynucleotides

Antisense RNAs, which are polynucleotides with base se-quences complementary to messenger RNAs, have beenfound in both prokaryotes and eukaryotes. Natural anti-sense RNAs are not common but synthetic RNAs directedat specific targets have been widely studied. It has beendemonstrated that they can function as inhibitors of mes-

senger RNA translation. In prokaryotes, the most effectiveinhibitors appear to have a base sequence complementaryto the 5′ leader region, including the Shine–Dalgarno se-quence, which is involved in the binding of mRNA tothe small ribosomal subunit. In eukaryotes, translation ofmRNA is inhibited by polyribonucleotides complemen-tary to the 5′ untranslated region of mRNA, indicating a di-rect effect on initiation as in prokaryotes. Polynucleotidescomplementary to the 3′ untranslated region of mRNAalso inhibit translation in some cells, and this effect maybe due to destabilization of mRNA by ribonucleases spe-cific for double-stranded RNA. The effect of antisensepolynucleotides is not restricted to translation of mRNAs,but transcription as well as the processing of transcriptsmay also be inhibited.

F. Availability of Amino Acids, tRNAAbundance, and Codon Usage

1. Amino Acids

Polypeptide synthesis depends on an adequate supplyof tRNAs charged with the 20 protein amino acids andappropriate interactions between their anticodons and thecodons of mRNA. Peptide chain elongation is decreased orinhibited by lack of amino acids or other conditions givingrise to an imbalance or deficiency in aminoacyl-tRNAs.

2. Abundance of tRNAs and Codon Usage

Different tRNAs are present in the cytosol in unequalamounts, and elongation rates are slower at codons corre-sponding to rare tRNA species.

The existence of synonymous codons raises the ques-tion of preferential use of some codons and its possible sig-nificance in relation to translational efficiency and control.In some bacteria (e.g., Pseudomonas aeruginosa, whichhas a high content of G + C, 67.2%, in DNA), the mostcommon codons are those with the strongest predictedcodon–anticodon interaction—that is, G + C base pairs—but this preference is not universal and, for example, doesnot apply to E. coli, which has a lower proportion of G + C(50%). Although codon usage may play a part in deter-mining elongation rates, it is probably of less importancein translational control than the secondary structure ofmRNA in relation to the rate of initiation of protein syn-thesis.

G. Modulation of Ribosome Activity

Specific ribosomal components have an important func-tion in relation to the fidelity of protein synthesis. Thus,in E. coli ribosomal protein S12 determines the accu-racy of codon–anticodon interactions and modulates the

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translational error frequency in the presence of the antibi-otic streptomycin.

To what extent reversible modifications of ribosomalconstituents are involved in translational control of pro-tein synthesis is uncertain. Although phosphorylation ofribosomal protein S6 increases with cell proliferation, itis not known whether this change is directly related to theaccompanying increase in protein synthesis by an effecton the translation rate.

H. Ribosome-Inactivating Proteins

Many molds and plants produce toxins, which are pro-tective reagents, termed ribosome-inactivating proteins(RIPs), directed at particular cells and their ribosomes.These toxins are classified as either type I or type II RIPsaccording to the number of polypeptide chains.

Type I RIPs comprise a single polypeptide chain; forexample, α-sarcin, an extracellular cytotoxin produced byAspergillus giganteus, consists of a single chain of 150amino acid residues.

Type II RIPs comprise two polypeptide chains, A andB. The A chain has the ability to inactivate ribosomes, andthe B chain is a galactose-specific lectin responsible for theentry of the toxin into the target cell. Ricin, which is iso-lated from castor beans, is representative of type II RIPs.

Ribosomes are inactivated as a result of the RNAN-glycosidase activity of RIPs. These toxins havedifferent specificities for particular cells and ribosomes.However, the target site for all RIPs is an adenylateresidue (position 2660 in the E. coli 23S rRNA sequence)located within a highly conserved sequence of 12nucleotides (5′

2654AGUACGAGAGGA26653′). Cleavageof the GpA2660 internucleotide bond or depurination ofA2660 is sufficient to inactivate the ribosome. The targetresidue is located in the loop region of a stem–loopelement of secondary structure which is termed theα-sarcin stem–loop. Thus, the target adenylate is eitherdirectly or indirectly essential for ribosome function. Theα-sarcin stem–loop is known to be important for bindingelongation factors EF-Tu and EF-G to the ribosome. RIPshave attracted interest as active components of reagentsdirected at particular targets such as cancer cells.


The control of protein synthesis, either by regulation ofthe amount of mRNA available for translation or by theefficiency with which it is translated, is important in cellgrowth and development as a factor determining the levelof cellular and extracellular proteins. Subversion of thiscontrol occurs in cells infected by viruses when the viral

nucleic acid uses the protein-synthesizing machinery ofthe host cell and thereby changes normal cell metabolismin favor of the synthesis of viral proteins needed for theproduction of virus progeny.

The polypeptide chains of all proteins are synthesizedby the process described above. This mechanism givesrise to primary polypeptide chains, which are often furthermodified—for example, by cleavage into smaller peptides,by structural modification of selected amino acid residues,by splicing of the polypeptide chain, or by the formationof covalent bonds between polypeptide chains. Some ofthese secondary modifications are related to the correctfolding of polypeptide chains and to the production ofactive enzymes or peptide hormones from inactive pre-cursors (e.g., insulin from proinsulin). Also, the transportof proteins within the cell or the secretion of extracellularproteins is often linked to structural changes in polypep-tide chains either during or after completion of synthesis.







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