Methods 56 (2012) 33–43

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Methods

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Review Article

Isolation, electron microscopy and 3D reconstruction of invertebratemuscle myofilaments

Roger Craig ⇑Department of Cell Biology, University of Massachusetts Medical School, 55 Lake Avenue North, Worcester, MA 01655, USA

a r t i c l e i n f o a b s t r a c t

Article history:Available online 2 December 2011

Keywords:Myosin filamentActin filamentThree-dimensional reconstructionStructureMethodologyInvertebrate

1046-2023/$ - see front matter � 2011 Elsevier Inc. Adoi:10.1016/j.ymeth.2011.11.007

Abbreviations: EM, electron microscopy; 3D, three-Helical Real Space Reconstruction; MLCK, myosin lightglycol bis(b-aminoethyl ether) N,N,N0 ,N0-tetraacetic a⇑ Fax: +1 508 856 6361.

E-mail address: roger.craig@umassmed.edu

Understanding the molecular mechanism of muscle contraction and its regulation has been greatlyinfluenced and aided by studies of myofilament structure in invertebrate muscles. Invertebrates areeasily obtained and cover a broad spectrum of species and functional specializations. The thick (myo-sin-containing) filaments from some invertebrates are especially stable and simple in structure and thusmuch more amenable to structural analysis than those of vertebrates. Comparative studies of inverte-brate filaments by electron microscopy and image processing have provided important generalizationsof muscle molecular structure and function. This article reviews methods for preparing thick and thinfilaments from invertebrate muscle, for imaging filaments by electron microscopy, and for determiningtheir three dimensional structure by image processing. It also highlights some of the key insights intofilament function that have come from these studies.

� 2011 Elsevier Inc. All rights reserved.

1. Introduction tion of muscle contraction. The major techniques used in under-

All muscles contract by sliding of the thick (myosin-containing)filaments past the thin (actin-containing) filaments (for overviewssee [1–3]). Essential to an understanding of contraction and its reg-ulation is knowledge of the molecular structure of the myofila-ments and the changes in their structure that occur in differentstates. Our anthropocentric view of biology and focus on the rele-vance of basic science to human disease means that most musclestudies have been carried out on vertebrates. However, some ofour greatest insights into muscle structure and function have comefrom invertebrate studies. For example, the greater stability ofinvertebrate thick filaments compared with their vertebrate coun-terparts has made possible more detailed structural analysis inthe invertebrates. The insights gained from these studies have pro-vided an essential foundation for understanding vertebrate fila-ment structure. In addition, invertebrates show a range ofspecializations in filament structure and organization related totheir specific needs. This range has made it possible to derive usefulgeneralizations concerning filament structure and function [2,3].Several model systems, including spiders, horseshoe crabs, insectsand mollusks, have played key roles in these advances. These sys-tems have provided fundamental insights into both thick and thinfilament structure, and the relation of this structure to the regula-

ll rights reserved.

dimensional; IHRSR, Iterativechain kinase; EGTA, ethylene

cid; UA, uranyl acetate.

standing filament structure include X-ray crystallography, X-rayfiber diffraction, spectroscopic methods such as electron paramag-netic resonance, and electron microscopy (EM). In this article I focuson electron microscopy and image processing techniques that leadto an understanding of three-dimensional (3D) filament structure.

2. Filament preparation

The thick and thin filaments that make up striated muscle arearranged in overlapping arrays within contractile units known asmyofibrils [1–3]. The state of muscle required to separate thickand thin filaments from each other, and to isolate them biochem-ically, is the relaxed state, in which filament interaction is minimal.The key step in isolating either thick or thin filaments is thereforehomogenization of muscle in a relaxing medium, whose criticalrequirements are the presence of Mg�ATP and the absence ofCa2+. A typical relaxing solution contains 0.1 M salt, Mg�ATP (todissociate actin from myosin), additional Mg2+ to stabilize the thinfilaments, and ethylene glycol bis(b-aminoethyl ether) N,N,N0,N0-tetraacetic acid (EGTA) to chelate Ca2+; it is buffered to pH �7.0[4–7]. We also add NaN3 to inhibit bacterial growth. Our normalsolution is: 0.1 M NaCI, 3 mM MgCl2, 5 mM Mg�ATP, 1 mM EGTA,3 mM NaN3, 10 mM sodium phosphate, pH 7.0 [4].

2.1. Native thick filaments

In a typical preparation (Fig. 1; [4–7]), muscle fiber bundles1–2 mm in diameter are removed from the freshly dissected animal

34 R. Craig / Methods 56 (2012) 33–43

and held at about rest length by securing to a frame with thread orby maintaining attachment to the rigid exoskeleton. The muscle isthen placed in cold relaxing solution. Two hundred milligrams(wet weight) of muscle is finely chopped with a scalpel, the piecesplaced in 3.0 ml ice-cold relaxing solution in a 15 ml tube, and thesuspension homogenized briefly with cooling on ice. Typically wegive 2 bursts each of 5 s at setting 5 on a Polytron homogenizer fit-ted with a 12 mm diameter probe. The exact time and setting de-pends on the muscle and type of homogenizer, and is determinedby trial and error; homogenization should be the minimum re-quired to release sufficient filaments, as excess homogenizationcauses filament breakage and denaturation. The homogenate iscentrifuged for 2 min at 15000g in a bench centrifuge at 4 �C to re-move large debris, leaving the filaments in the supernatant. At thisstage the suspension is at fairly high concentration and appearsmilky. This crude suspension contains both thick and thin filamentsas well as other cellular components such as fragmented organelles.For electron microscopy, the suspension is diluted 5 to 10-fold withrelaxing solution to provide a good distribution of filaments on theEM grid without too much filament overlap; the dilution factor isdetermined by trial and error.

A variation on this method that is often used [4–6] is to firstchemically demembranate the muscle (held at rest length) bytreating with detergent (e.g. 0.1% w/v saponin) in relaxing solutionfor �3 h at 4 �C with agitation. This allows soluble protein to exitfrom the fibers, resulting in a cleaner filament preparation withless background protein. Use of relaxing solution maintains the fil-aments in a relaxing environment. The muscle is then chopped andhomogenized as above.

If it is desired to purify the thick filaments by removing the thinfilaments (for example for cleaner EM images, or to determine fila-ment constituents by SDS–PAGE), further differential centrifugationwould seem the obvious choice, due to the larger S-value of thickcompared with thin filaments. In practice this works poorly, and sed-imented thick filaments are invariably accompanied by many thin fil-aments. This appears most likely to be due to the trapping of thinfilaments in the sedimenting mesh of thick filaments, or possibly tothe presence of weak actin–myosin interaction even in relaxing con-ditions [8]. A simple solution to this problem has been devised, usingthe actin-severing protein gelsolin to cut the thin filaments into shortsegments, which are not readily enmeshed and which interact lessreadily with the thick filaments [8]. Gelsolin normally requires thepresence of micromolar Ca2+ for activity, which is incompatible withthe relaxing conditions required to keep filaments separated. Ca2+-insensitive gelsolin (the N-terminal half) can be prepared from plas-ma gelsolin by enzymatic digestion with thermolysin [8], or can bebacterially expressed [9]. Both methods yield a molecule that seversthin filaments into short segments under relaxing conditions. Fol-lowing gelsolin treatment, the suspension is centrifuged (18,700gfor 15 min) to produce a pellet containing relatively pure thick fila-ments [8]. Re-suspension and re-centrifugation in relaxing solutioncan be carried out to further purify the filaments (Fig. 3 cf. Fig. 2a). Inaddition to treating filaments in suspension, gelsolin can also beused simply to rinse an EM grid with adherent filaments. The thin fil-aments rapidly disappear.

Fig. 1. Schematic of main steps in

2.2. Native thin filaments

While thick filaments from different invertebrate species havevarying helical parameters (and are thus useful for comparativestudies; [3,10,11]), thin filaments from all muscles (both vertebrateand invertebrate) are remarkably similar in structure, and studiesof invertebrate thin filaments have therefore played a less impor-tant role in our understanding of muscle molecular structure. Anexception is a negative stain EM study of Limulus thin filaments,which was the first to directly visualize tropomyosin movementon actin that is the basis of thin filament regulation (see Sec-tion 4.2.2) [12]. This strongly supported earlier modeling studiesof X-ray diffraction data [13–15]. Limulus filaments were chosenfor this study because they have a larger (therefore more visible)troponin than vertebrates and exhibit excellent Ca2+-sensitivityof actin-activated myosin ATPase. They were therefore deemedmost likely to reveal any changes in thin filament structure in re-sponse to Ca2+.

Thin filaments from Limulus and other invertebrates are pre-pared as described by [16] with modifications by [17] and [18].Muscle fiber bundles are tied to sticks at close to rest length, trea-ted overnight with 50% glycerol/water to permeabilize the cellmembranes, teased into millimeter bundles while still attachedto the sticks, and placed in 0.1 M KCl, 5 mM phosphate, 3 mMMgCl2, 3 mM EDTA (ethylenediamine-N,N,N0,N0-tetraacetic acid),pH 7.0 for 10 min. The fiber bundles are then homogenized in thissolution. The rigor myofibrils thus produced are collected by cen-trifugation (5 min at 5000g), re-dispersed manually, and washedthree times by centrifugation and resuspension. The myofibrilsare then resuspended in a low salt relaxing solution (20 mM KCl,0.1 mM EDTA, 1 mM MgCl2, 5 mM ATP, 5 mM phosphate, pH 6.0),homogenized to dissociate the thick and thin filaments, and centri-fuged (5 min at 5,000g) to remove coarse myofibrillar fragments.The supernatant is then centrifuged more strongly (1 h at25,000g followed by 30 min at 50,000g) to sediment the thick fila-ments. Although many thin filaments will also sediment, some areleft in the supernatant, and these can be collected by sedimenta-tion at 100,000g for 3 h. The pelleted filaments are resuspendedand used for grid preparation.

3. EM observation

Filaments prepared as above can be studied by a variety of elec-tron microscopic methods. These include negative staining, metalshadowing and cryo-electron microscopy.

3.1. Negative staining

3.1.1. PrincipleNegative staining is a technique by which macromolecular

assemblies such as filaments can be easily studied at moderatelyhigh resolution and with good contrast and structural preservation.Stain surrounds and supports the structure and penetrates itsinterstices, revealing detailed surface shape at resolutionsapproaching 2 nm. A drop of filament suspension is applied to a

preparation of thick filaments.

Fig. 2. Negative staining and shadowing. (a) Unpurified tarantula thick filaments,with thin filaments in background (from [4]); (b) metal-shadowed tarantula thickfilament showing right-handed helical tracks (from [5] with permission); (c)negatively stained thin filament from frog cardiac muscle (from [103] withpermission); this vertebrate image is chosen for the clarity with which it revealstropomyosin (arrowed strands), although it was invertebrate filaments that firstdemonstrated the tropomyosin-shift mechanism unequivocally [12]. In (a and c),protein is white and stain dark; in (b) metal is white. Scale bars: a, b (100 nm), c(50 nm).

R. Craig / Methods 56 (2012) 33–43 35

carbon-coated EM grid, followed by several drops of stain. The lastdrop is removed with filter paper and the remaining stain allowedto dry. The grid is then ready to image in the EM. The best negativestain for thick and thin filament observations is aqueous uranylacetate (UA), generally 1–2% w/v. While UA also gives a degree ofpositive staining, its key advantage is its high contrast and the factthat it also acts as a rapid fixative [19]. Thus, when the stain dries,and its concentration increases, filament structure is not disruptedby the increased ionic strength. This is a key point, as thick fila-ments dissolve at high salt levels. The final thickness of the driednegative stain is an important factor in obtaining good images.Optimally the stain should be similar in thickness to the filamentbeing studied, so that the whole depth of the filament is con-trasted. All thin filaments are about 10 nm in diameter and requirea relatively thin layer of stain. Thick filaments vary (depending onspecies) generally in the range 30–40 nm, and are best contrastedwith a thicker layer of stain. This can be achieved by leaving aslightly thicker layer of the liquid stain on the grid before dryingand/or by using a higher concentration of stain.

3.1.2. UsesNegative staining of invertebrate thick and thin filaments has

played a key role in our basic understanding of myofilament struc-ture and function (Fig. 2). It was the first technique to reveal thebipolar organization of myosin molecules in thick filaments [20],and the 3D organization of actin in thin filaments [20,21]. Nega-tively stained Limulus thick filaments were the first in which the3D helical organization of myosin heads was preserved, makingpossible the first 3D reconstruction of a thick filament [22,23].Studies of other invertebrates, together with improved stainingand 3D image processing techniques, yielded additional insightsinto thick filament structure (Fig. 2a) [5,6,24,25]. Negative staininghas also provided key insights into the structure of thick filamentsin invertebrate smooth muscles [26–28]. In addition, the techniquehas revealed the structural changes that occur in invertebrate thick

filaments upon activation by Ca2+ binding or myosin phosphoryla-tion [4,29,30]. Such routine negative staining techniques have beenfurther developed to enable the capture of transient structuralchanges occurring on the millisecond timescale [19]. This time-re-solved technique reveals the changes in scallop thick filamentstructure that occur in the first 10–30 ms after Ca2+ binding [31].In the case of thin filaments, comparative studies of negativelystained invertebrate filaments revealed the common helical orga-nization of actin present in diverse muscles [21], and it was nega-tive staining of Limulus filaments in the presence and absence ofCa2+ [12,32] that first visualized the steric blocking model of thinfilament regulation originally postulated on the basis of X-ray dif-fraction data [13–15] (see Section 2.2 and 4.2.2; Figs. 2c and 5a–d).

3.1.3. ProcedureTechnical details, and the many variations on negative staining

technique applied to biological samples, can be found in [33,34].Here the approaches used for studying muscle filaments in our lab-oratory are outlined. EM grids, generally 400 (or finer) mesh, arecoated with carbon to act as a support for the filaments. This is aroutine technique available in any biological EM lab. Carbon is firstevaporated onto freshly cleaved mica in a vacuum evaporator, thenfloated from the mica onto a water surface. The floating carbon ispicked up by lowering the water surface onto grids laid out on awire mesh under the water, or by bringing individual grids heldin forceps through the water surface to pick up the carbon from be-low. Image contrast is best when the carbon layer is thin, improv-ing the signal over the background noise; however, thin carbon isfragile and easily breaks before or during observation under theelectron beam, unless the support grid mesh is extremely fine.One way to achieve this is to first coat grids with a thick (thereforestrong) ‘‘holey’’ carbon film, and use this as an extremely fine mesh(see Appendix for holey film method). A very thin layer of carbonfrom mica can be picked up on such a grid and is stable over thenarrow span of the holes (1–5 lm, compared with an �40 lmopening in a 400 mesh grid). Filaments are imaged over the thincarbon on the holes. Structural detail is even better when thereis no carbon underlying the filaments at all [20,35]. This can beachieved by applying filaments to a holey carbon film withoutany carbon over the holes. Filaments often remain in the film ofstain stretching across the hole. However, this film is extremelyfragile and usually breaks during observation in the electron beam(if not before), squashing or stretching the filaments. It can be sta-bilized by applying an extremely thin layer of carbon to the gridafter staining [36]. Apart from improved contrast and lower back-ground noise, an advantage of this method is that the top and bot-tom of the filament are more equally stained. A disadvantage ismore work.

The suspension of thick or thin filaments is applied to thecoated grid in a 5 ll droplet, which is allowed to rest on the gridfor about 30 s. The excess droplet is wicked off from the edge ofthe grid with filter paper (making sure that the thin layer left doesnot dry out!) and then several (6–10) drops of stain are run acrossthe grid. The last one is also wicked off and the grid allowed to dry.To obtain optimal staining and structural detail, an appropriate fil-ament concentration is required. Too many filaments and the staintends to form dense patches and many filaments overlap. Too fewand it is difficult to obtain the requisite good spreading of the stain.The optimal load is determined by trial and error, but a good start-ing concentration for both thick and thin filaments is about 1 lM.

3.1.4. Problems and fixesThe most common problem with negative staining of biological

samples is ensuring that the stain spreads to a thin layer that sur-rounds the structure. Very frequently during drying, the stain pullsaway from the grid surface to form a tiny droplet in one small area,

36 R. Craig / Methods 56 (2012) 33–43

due to the high surface tension of the aqueous stain and the hydro-phobic nature of most carbon films. The result is poor filamentstructure and contrast (often the filament will appear positivelystained). This problem can be avoided or minimized in severalways: one is glow discharge or UV treatment of the coated gridsto make them hydrophilic [37]. We have also found that slow dry-ing of the grid in a humidity chamber at about 80% relative humid-ity, or alternatively very rapid drying under a hair dryer can be veryhelpful. Because spreading of stain is aided by high humidity, labsin more humid environments have less trouble with negativestaining than those in dry conditions (e.g. winter in the northernUS).

A few special tricks have been developed that help with specificproblems. To preserve thick filament structure optimally, it hasbeen found that treatment with tannic acid before UA is useful[38,39]. Filaments are usually partially flattened during drying ofthe negative stain. This is lessened if (as mentioned above) theamount of stain used is such that the dried layer is roughly thesame thickness as the filament (any thinner, and support is lessand only part of the thickness may be contrasted; thicker, and con-trast is reduced and noise added due to excess stain). Flattening isalso reduced with tannic acid [39] or by including a trace of glyc-erol in the stain [40].

In the case of thick filaments, it is found that the helical order-ing of the myosin heads, known to be present in the relaxed state,is easily disrupted. This precludes 3D image reconstruction (Sec-tion 4). There are several causes. One is the binding of the heads(which are flexibly attached to the filament) to the carbon sub-strate; this is worse with glow discharged or UV-treated gridswhich produce a charged carbon surface that attracts the heads[41]. Thus surface treatment to ensure spreading of stain mustnot be overdone, and the use of high humidity or rapid dryingmay be preferred over glow discharge or UV treatment of grids. An-other, less obvious issue is the finding that UA staining of thick fil-aments that have been prepared in a chloride-based relaxingsolution usually causes disordering of the heads. The reason isnot known; Cl� is known to bind strongly to myosin filaments[42], but is present naturally inside cells only at low concentrations(�4 mM). This Cl� effect can be avoided by rinsing the filaments onthe grid with a solution containing acetate in place of Cl� [5,7].Thick filament helical order can sometimes be improved at lowerpH (6.5 rather than 7 or 7.5) and at lower ionic strength, althoughat the expense of filament bundling. The most stable ordering ofthe heads is achieved in the presence of the myosin inhibitorblebbistatin [43]. In filaments regulated by myosin RLC phosphor-ylation, phosphorylation causes disordering of the heads [4]. Im-proved order should therefore also be obtained in the presence ofkinase inhibitors or of phosphatase, although we have not yetdetermined whether this is actually the case. Making grids as soonas possible after filament preparation should also minimize un-wanted phosphorylation.

Finally, the cleanest images of thick filaments (best suited forimage reconstruction) are obtained if thin filaments are removed,so that filament superposition is minimized. While this is notessential (good images can be obtained by sufficiently diluting fil-aments), it is advantageous (see Section 2.1).

In the case of thin filaments, some of the best staining is ob-tained in the presence of phosphate ions (e.g. 5 mM) [35]. Againthe reason is unclear. Phosphate (as the inorganic ion or as ATP)is often present in solutions either as a buffer (Pi) or in relaxingsolutions (Mg�ATP). However, electron microscopists tend to sub-stitute other buffers for Pi when negative staining with UA becausethe stain rapidly forms an insoluble precipitate with Pi, makinggrids dirty. This problem is easily overcome, so that one can takeadvantage of the UA/Pi combination, simply by ensuring that thetwo solutions do not mix for a significant length of time. If the grid,

with filaments attached in a Pi-containing buffer, is rapidly flushedwith 8–10 drops of UA, precipitation is not a problem [35].

3.1.5. ImagingFollowing preparation, grids can be immediately observed in

the microscope, or they can be stored for weeks to months withoutsignificant deterioration. Images can be obtained by routine meth-ods if fine details of structure are not critical. The grid is usuallysearched at low magnification (2000–3000�) to find areas ofappropriate filament distribution and good negative staining (stainquality can vary a lot over the grid), then imaged at high magnifi-cation (30,000–60,000�). If 3D reconstruction is desired and imagedetail is critical, then low dose imaging methods are preferable.Low dose microscopy minimizes damage to the specimen by elec-tron irradiation. Typically electron micrographs are recorded afterfirst focusing on the area of interest. This requires a relativelybright beam and several seconds of irradiation, during which timethe filament will have been exposed to many thousands of elec-trons per square nm. Such irradiation causes the stain to migratefrom its original fine distribution, to form larger pools that no long-er delineate the finest details [44]. These details are best preservedif focusing is carried out to one side of the area to be imaged, sothat the actual image recorded receives minimal electron doses<1000 electrons/nm2 [5]. Low dose software making this possibleis available on many modern microscopes.

3.2. Shadowing

Filaments can also be imaged by metal shadowing techniques.While shadowing is usually less useful for understanding detailedstructure, it has proved critical in determining one aspect of myo-filament structure: the ‘‘handedness’’ (left or right) of the filamenthelix. Negative staining provides a projection image in which thetop side and underside of the filament are superimposed. The handof the helix is therefore unclear because the underside of a right-handed helix resembles the top side of a left-handed helix and viceversa. This issue is overcome by shadowing because only the topside of the filament is contrasted. In this way it has been shownthat all myosin and actin filaments so far studied have long-pitchhelices that are right-handed [5,6,45,46] (Fig. 2b). Shadowing hasalso been ingeniously employed to demonstrate myosin heads onthe surface of invertebrate smooth muscle thick filaments (nor-mally difficult to demonstrate by negative staining alone) [26,47].

Shadowing of thick and thin filaments can be carried out as fol-lows. A drop of filament suspension is applied to a carbon-coatedEM grid as for negative staining (mica, normally used for singlemolecule shadowing, is not desirable as it disrupts the helical orga-nization of myosin heads). The filaments are fixed briefly by rinsingwith 1% aqueous UA, rinsed with water then 30% (v/v) glycerol inwater, and the excess wicked off to leave a thin film [6,46]. Thesesteps help to strengthen the filaments against drying forces thatwould otherwise flatten them in the next step. The grid is thenplaced in a vacuum evaporator, and the chamber evacuated. Heavymetal (Pt) is evaporated onto the filaments at an angle of �12–30�.Following shadowing, grids are observed in the EM. Low dose tech-niques are unnecessary here due to the robustness of the metalcoating.

3.3. Cryo-EM

3.3.1. PrincipleCryo-EM is a method for observing macromolecular structures

without fixation, staining or drying. ‘‘Frozen-hydrated’’ specimensare observed at low temperature in thin aqueous films suspendedover holes in a carbon support film. Cryo-EM is thus free of the dry-ing, staining and fixation artifacts present in other techniques. It

R. Craig / Methods 56 (2012) 33–43 37

provides the closest to in vivo filament images of biological struc-tures available, although it comes with its own attendant difficulties.

3.3.2. UsesEarly studies showed that the helical order of invertebrate myo-

sin filaments could be well preserved by cryo-EM [48], thus avoid-ing artifacts of negative staining. However, 3D reconstruction wasnot carried out. The first demonstration that thick filaments couldbe reconstructed from cryo-EM images was with scallop filaments[49], although head conformation could not be interpreted in detailin this work. This problem was overcome in studies of tarantula fil-aments, which have highly ordered head arrays: in this case cryo-EM played a crucial role in deciphering the organization of myosinheads, which had proved elusive for over twenty years using neg-ative staining (Fig. 3; see Section 4.3.2 and 4.4) [11,50]. Subsequentstudies have shown that similar head arrangements are present ina variety of other invertebrates (Limulus [51], scallop [52,53]) andalso vertebrates (mouse cardiac muscle [54]). While the earliernegative staining studies had preserved the helical organizationof the heads, their relatively low resolution (5 nm) left the exacthead arrangement ambiguous [55]. Most interpretations suggestedthat the heads of each myosin molecule were splayed apart fromeach other, extending up and down the filament to the next levelin each direction [5,24,56]. Cryo-EM combined with single particleimage processing showed unambiguously that this model waswrong: the two myosin heads from each molecule in fact interactwith each other intramolecularly in a compact structure, in away that immediately suggests how this might function to switchactivity off in relaxed muscle (Section 4.3.2) [50]. Cryo-EM has alsoplayed a key role in our current understanding of thin filamentstructure and regulation [57–60], although this work was not oninvertebrate filaments. An invertebrate filament structure thathas been elucidated by cryo-EM is that of arthrin, a mono-ubiqui-tinated form of actin found in some insect flight muscles [61].

3.3.3. ProcedureFor cryo-EM, holey carbon films are generally used, as described

earlier for negative staining (true holes are used, without thin car-bon on top). The goal is to suspend the filaments in solution so that

Fig. 3. Cryo-EM image of purified, frozen-hydrated relaxed thick filaments fromtarantula muscle. Thin filaments have been removed by treatment with gelsolin,creating a cleaner image (cf. Fig. 2a). Bar = 100 nm. From [50].

forces that might affect their structure (e.g. by binding to a carbonsurface) are avoided or minimized. The holey films can be home-made (as described for negative staining) or purchased (Quantifoil(SPI Supplies, West Chester, PA) or C-flat (EMS, Hatfield, PA)). Quan-tifoil and C-flat grids have a regular array of holes of defined sizerather than the random size and location in home-made grids –but the cost exceeds $5/grid. Specimens are prepared as follows (de-tails can be found in [62]). A droplet of filament suspension (5 ll) isapplied to a grid as for negative staining. The grid is held in forceps ina plunging device, ready to be dropped into liquid ethane (cooled byliquid nitrogen), to cause vitrification (freezing without ice crystalformation). After 30 s, the grid is blotted gently with filter paper(from one or both sides) to form a thin film. The grid is immediatelyplunged into the liquid ethane to freeze the thin film before it dries.The grid is then transferred to a pre-cooled grid box under liquidnitrogen and stored in a liquid nitrogen Dewar until ready for imag-ing. Liquid nitrogen and liquid ethane are both cryogens and must behandled with care. Liquid nitrogen can cause skin burns, and if usedin a totally sealed room could cause suffocation. Liquid ethane alsocauses burns, and in the gaseous form is potentially explosive. It istherefore used in small amounts (�1–2 ml), and is disposed of byallowing it to evaporate in a fume hood.

Grids for cryo-EM are first glow discharged in an atmosphere ofamylamine within 2 h of use, to make the surface hydrophilic. Thisensures that when the grid is blotted, the thin aqueous film re-mains spread. For muscle filaments and other structures that aresalt-sensitive, blotting must be performed in a humid atmospherein order to minimize evaporation between the times of blottingand freezing. Evaporation would otherwise lead to disruptive in-creases in ionic strength (high ionic strength for example, causesmyosin heads to disorder [63] and filaments to dissolve). A Plexi-glas chamber supplied with humid air from a domestic humidifierworks well, or commercial, humidity-controlled plunge-freezingdevices are available [64]. These devices, under computer control(Gatan (Pleasanton, CA) model CP3; or FEI (Hillsboro, OR) Vitrobot),are capable of giving reproducible ice thickness and humidity con-trol, but are expensive (tens of thousands of dollars).

3.3.4. ImagingFrozen grids are placed in a pre-cooled (to approximately liquid

nitrogen temperature) cryo specimen holder (e.g. Gatan 626) in acryo-transfer station. The holder is then inserted into the microscopewhile observing precautions to avoid ice contamination on the gridsurface (by having the grid covered with a movable shutter duringtransfer, and carrying out the transfer operation quickly). The holderis left in the microscope for about 30 min to allow the vacuum and tem-perature to stabilize. The shutter is then opened and the grid searchedat low magnification (�2000�) for areas of suitable ice thickness.Images are acquired under low dose conditions (<1000 electrons/nm2), which are mandatory for cryo-EM, as aqueous specimens inthe absence of stain are rapidly destroyed in the electron beam.

3.3.5. IssuesThe key advantage of cryo-EM is its ability to capture and image

close-to in vivo macromolecular structure. In addition, the entirestructure (including the interior) is visualized, not just the surface(as with negative staining and shadowing). Cryo-EM has trans-formed our understanding of thick filament structure, and inverte-brates have been essential to this success [50]; cryo-EM has alsosucceeded at imaging F-actin filaments at high resolution [59,60].However, cryo-EM comes at a cost. It is slower, technically more dif-ficult, and requires more advanced instrumentation. Image contrastis low due to the absence of stain. Images must therefore be recordedat higher defocus (�1.5–3.0 lm) than with stained specimens inorder to generate some phase contrast – but this compromisesresolution. Many grids are useless because the ice is too thick or

38 R. Craig / Methods 56 (2012) 33–43

too thin. Low dose imaging means that the operator is ‘‘shootingblind’’, so that many images are later found to contain few or no fil-aments. When using film, the operator does not know whether suc-cess has been achieved until the film is processed: the advent of CCDcameras has diminished this difficulty greatly as images can beviewed immediately after acquisition, providing immediate feed-back on specimen quality.

3.3.6. Choosing between negative stain and cryo-EMBoth techniques can yield high resolution insights into muscle fil-

ament structure. The key advantage of cryo-EM is the imaging ofstructure in its native form. However, negative stain is sufficientfor answering most questions, and the added difficulty of cryo-EMmeans that in many cases, negative stain is to be preferred [65]. Itis well established that in most instances, negative stain providesexcellent fidelity of structural preservation, so that it can be usedto answer moderate resolution questions. An example – negativestain has been used in many papers to study the effects of mutationsin tropomyosin, actin or troponin on the regulatory movements oftropomyosin (e.g. [66,67]). Carrying out the same experiments usingcryo-EM would have taken years longer. Any potential additional in-sights may not have justified the greater effort.

3.4. Sectioning techniques: conventional and cryo-sectioning

3.4.1. PrincipleIf one is interested in understanding filament structure and

interactions in the context of the filament lattice of the sarcomere,sectioning techniques are required. Muscle specimens (live orchemically demembranated) are fixed, embedded, and sectionedeither transversely or longitudinally. Stained sections are observedin the microscope, and the structure, arrangement and interactionsof filaments can be studied in this way.

Fig. 4. (a) Longitudinal section of rigor insect flight muscle, showing alternating thick andaverage of one chevron-like structure similar to those in (a) calculated from tomogram o(those closer to and further from the pointed end of the thin filament, respectively). The cpermission. Scale bar = 100 nm.

3.4.2. UsesRoutine fixation (glutaraldehyde followed by osmium tetrox-

ide) and routine embedding/sectioning (epoxy resin, room tem-perature sectioning) methods have been applied to bothvertebrate and invertebrate muscles many times, leading to ourknowledge of basic filament and sarcomere structure [2]. Oneinvertebrate muscle stands out in going well beyond this basicinformation: asynchronous insect flight muscle. The quasi-crystal-line organization of filaments in this muscle has led to critical in-sights into the interactions between actin and myosin in differentphysiological states – insights applicable to understanding themolecular basis of contraction in all muscles (Fig. 4a) [68,69].Why is asynchronous insect flight muscle so useful? The answerlies primarily in its filament organization. In these muscles, usedto generate wing movements, the sarcomere is organized withthin filaments in the same plane as nearest neighbor thick fila-ments, and all thick filaments in rotational register. Thus verythin longitudinal sections (�20 nm) can be cut, containing a sin-gle layer of actin and myosin filaments (myac layers), and theimages are especially useful for image analysis and averaging.This contrasts with vertebrate and most other muscles (wherethin filaments are not in the same plane as nearest neighbor thickfilaments, and thick filaments can have varying rotations). Thespecies most frequently used for studies of insect flight muscleis the giant water bug Lethocerus. Early studies of myac layersproduced beautiful images of myosin heads in relaxed and rigorstates, which led to some key concepts of the crossbridge cycleof contraction, including the idea that myosin heads wentthrough a large change in angle while attached to the thin fila-ment to generate the powerstroke [68]. While this idea has sincebeen modified, it provided a simple framework for thinking aboutthe contractile mechanism. More recent cryo studies of insectflight muscle have led to many other insights into the molecularbasis of contraction (see Section 3.4.4).

thin filaments with crossbridges joining them; M-line at top, protein black. (b) Classf rapidly frozen muscle, showing interpretation in terms of ‘‘lead’’ and ‘‘rear’’ headsentral filament is actin with partial myosin filaments on either side. After [72], with

R. Craig / Methods 56 (2012) 33–43 39

3.4.3. Conventional fixation and sectioningMuscles are used fresh or chemically skinned (with detergent or

glycerol), fixed with glutaraldehyde, postfixed with OsO4, stainedwith uranyl acetate, dehydrated with an ethanol or acetone series,embedded in epoxy (Araldite or Epon), then sectioned on a dia-mond knife [33,68]. The stunning contrast seen in many myacimages of insect flight muscle results from the (unconventional)use of potassium permanganate as a section stain [69]. Severalvaluable variations on this procedure have been developed, includ-ing the use of tannic acid as an additional fixative, which improvesboth contrast and preservation [70].

3.4.4. Rapid freezing/freeze substitutionAlthough these routine methods are very powerful, they are

limited in the level and fidelity of molecular detail they can pro-vide. In general, crosslinking by glutaraldehyde appears to disruptthe helical organization of myosin heads in thick filaments, whilemolecular details of the thin filaments are also lost. Low tempera-ture variants of these techniques applied to invertebrate muscleshave proved very successful at overcoming some of these limita-tions. A powerful procedure for improving molecular preservationin sections is the use of fast physical fixation (rapid freezing) ratherthan slow chemical fixation (chemical crosslinking). The muscle fi-ber bundle is mounted on a specimen disk attached to a plungingdevice which is dropped onto a flat, polished copper block cooledwith liquid helium [71,72]. The outermost 1–10 lm of the tissueare rapidly frozen (�1 ms) without damage due to ice crystal for-mation (immersion in liquid nitrogen, by comparison, is a slowmethod of freezing and results in unacceptable ice crystal damage).Following this rapid physical fixation, the specimen is freeze-substituted, a procedure carried out at ��80 �C in which the icein the specimen is slowly replaced by organic solvent (usually ace-tone) containing chemical fixative (OsO4 or tannic acid or UA, orsome combination) [71,72]. In this way chemical fixation takesplace at low temperature on tissue already stabilized by physicalfixation. The result is superior preservation, even of the labile myo-sin head helical organization. Following freeze-substitution, speci-mens are embedded and sectioned conventionally. Using thisapproach, it has been possible to demonstrate the helical organiza-tion of myosin heads in thick filaments in situ in tarantula muscleand to determine directly the rotational symmetry of the headorganization in both tarantula and scallop muscle by observingthick filaments in transverse section [73–75]. Similar detail hasalso been observed in vertebrate muscle [76–78] leading to im-proved insights into vertebrate sarcomeric structure. The moststriking use of this technique has been in detailed studies of insectflight muscle [72], in which three-dimensional information hasbeen obtained by electron tomography of sections and tomogramaveraging. This is described in Section 4.1.2.

3.4.5. Cryo-sectioningAn alternative cryo approach involves low temperature section-

ing (cryo-sectioning) followed by ambient temperature observa-tion. This has proved extremely powerful in the analysis of thickfilament structure in vertebrates [79]. In the case of invertebrates,detailed filament structure has also been observed. In one study,skinned muscles were put into ATP-free solution, creating strongrigor crosslinks between the myosin and actin filaments, then trea-ted with cryo-protectant and plunge-frozen in liquid ethane [80].Specimens were cryo-sectioned at �110 �C [81], the sections col-lected on a grid, thawed, and negatively stained with uranyl ace-tate. Because the filaments were held together by rigorcrosslinks, it was not necessary to fix before freezing. These sec-tions made it possible to observe filament molecular detail (with-out fixation artifacts) at the high resolution possible with negative

staining (see Section 3.1) but here in the context of the filamentlattice.

The most faithful representation of filament structure in the fil-ament lattice is obtained, as with isolated filaments, by cryo-EM. Inthis case, cryo-sectioning is carried out on unfixed, rapidly frozentissue, and sections are observed in the frozen-hydrated state[82,83]. As with all unstained specimens, contrast is low and imag-ing must be carried out by low dose techniques. Excellent nativefilament detail has been obtained from sections of insect flightmuscle [84], confirming inferences from more artifact-prone tech-niques involving fixation, staining or filament isolation. Cryo-sec-tioning of unfixed and un-cryoprotected tissue is difficult andcomes with its own artifacts [82,83].

4. Three-dimensional reconstruction

Owing to the large depth of field in the electron microscope,images obtained are two-dimensional projections of 3D structures.Top sides and under sides of filaments are thus superimposed andcannot be distinguished in a single image. There are several waysto obtain 3D filament information, all requiring different angularviews of the filament subunits. Three powerful 3D reconstructiontechniques commonly used are electron tomography and two ver-sions of helical reconstruction. These methods combined with thepreparative techniques described earlier have provided key in-sights into myofilament structure in invertebrates.

4.1. Electron tomography

4.1.1. PrincipleWith electron tomography, the specimen grid is tilted in the

electron microscope and the information obtained from multipletilt views is combined by back projection to create a 3D reconstruc-tion [72,85]. The microscope must be equipped with an accurate(preferably motorized and automated) goniometer tilt stage (com-mon on modern microscopes). Images are typically collected over atilt range of +/�60�, and typically at62� intervals (dependent uponthe thickness of the object being imaged and the desired resolu-tion). The physical limitations of holders and stages prevent imagesbeing obtained at much greater than 70� tilt, resulting in loss ofsome information in the Z direction (corresponding to the thick-ness of the filament). Modern microscopes equipped with CCDcameras and software that automates specimen tilting, focusingand image acquisition enable a full set of images to be collectedin an hour or so. Automation also makes possible a large decreasein the electron dose required to obtain a tilt series, resulting in im-proved resolution. Tomographic reconstruction programs areavailable at no cost (e.g. IMOD [86]) or are sold as a part of atomography package with modern microscopes.

4.1.2. UsesSome of the most impressive insights into actomyosin filament

interaction have come from electron tomography of sectioned in-sect flight muscle (used because of its high degree of order; seeSection 3.4.2). Many of these studies have used rapidly frozen,freeze substituted muscle, which not only preserves molecularstructure optimally (Section 3.4.4), but also makes possible thecapture of intermediate structural states of myosin crossbridgesduring contraction [72]. A key innovation in this work has beenthe computational classification of different structural motifs (rep-resenting, for example, myosin heads attached to actin in differentpatterns and conformations) and the 3D averaging of motifs withinclasses, enabling detailed structural information to be extractedfrom heterogeneous populations (Fig. 4b). The resolution of thesestudies is often of the order of 5 nm, and individual myosin heads

40 R. Craig / Methods 56 (2012) 33–43

attached to actin, as well as troponin molecules on the thin fila-ments, are thought to have been visualized and their conforma-tions elucidated by fitting of myosin head atomic structures (seeSection 4.4). These studies have led to insights into the effect ofadding different nucleotides to rigor muscle, the visualization ofa wide range of myosin head structures during isometric contrac-tion, and the ability to determine the effects of transient mechan-ical perturbations on myosin head conformation [72].

4.2. Helical reconstruction

4.2.1. PrincipleHelical structures, such as actin and myosin filaments, display

multiple views of their subunits in a single image, due to theintrinsic nature of a helix. If the symmetry of the helix (the axialrise and rotation per subunit) is known, then helical reconstructiontechniques, taking advantage of these views, can be used to com-pute a 3D structure from a single image [87–89]. Such reconstruc-tions reveal only the features in the structure that obey the helicalsymmetry. Individual reconstructions are typically noisy and varyone from another, for example due to variations in stain distribu-tion in negatively stained specimens. Individual reconstructionsare therefore averaged to create a reconstruction with a higher sig-nal to noise ratio with improved resolution. To carry out a helicalreconstruction, digitized electron micrograph images are computa-tionally boxed, then straightened, and their Fourier transforms cal-culated. By combining the indexing information of the transform (afunction of the helical symmetry) with the layer line data, a 3Dreconstruction (density map) can be computed. Multiple recon-structions are averaged by aligning and averaging the filament datain Fourier space then computing the reconstruction from the aver-aged data [89].

4.2.2. UsesHelical reconstruction techniques have played a key role in our

understanding of both thin and thick filament structures. Earlyreconstructions of F-actin were based on vertebrate proteins, but,as described earlier, it was helical reconstruction of negativelystained invertebrate filaments (Limulus) that first directly revealed[12,32] the physical blocking mechanism by which tropomyosinswitches off thin filaments, and the way in which Ca2+ acts to movetropomyosin, making previously blocked myosin binding sitesaccessible (Fig. 5a–d; Sections 2.2 and 3.1.2) [13–15]. Reconstruc-

Fig. 5. Three-dimensional reconstruction of thin and thick filaments. (a–d) Helical reProjections down long-pitch helices, showing outer and inner domains of actin subunits (and inner domain in high Ca2+ state (b). (c and d) Surface presentation of the same reconIHRSR reconstruction of tarantula thick filament based on cryo-EM images. The repeatiFig. 6). From [50]. Scale bar for (c–e) = 10 nm.

tions of thin filaments from Drosophila with engineered mutanttroponin have provided insights into critical residues involved inthe regulatory movement of tropomyosin [90].

Similarly, it was helical reconstruction (of negatively stainedLimulus thick filaments) that provided the first visualization ofthe arrangement of myosin heads on helical tracks in a thick fila-ment (Section 3.1.2) [22]. In work on vertebrates, the helical orderknown (from X-ray diffraction) to be present in intact muscle [91]was lost during filament isolation and staining [20,41]. Successwith Limulus was a result of the greater stability of the myosinhead arrays compared with vertebrate filaments, and the develop-ment of gentle methods of filament isolation that preserved myo-sin head order [22]. Helical reconstruction of Limulus filamentsdirectly revealed the 4-fold arrangement of myosin head helicaltracks on the filament surface [22]. Subsequent studies of scallop,scorpion, tarantula and insect filaments yielded similarly strikingreconstructions [5,6,24,25,49], and demonstrated some of thediversity of invertebrate filament structure. In the case of the clo-sely related Limulus, scorpion and tarantula [5,22,24], it was shownthat all filaments had a similar structure and 4-fold rotational sym-metry. In contrast, scallop was shown to have a 7-fold arrangementof heads [6]. In the initial studies [6,22] the two heads of each myo-sin molecule were not resolved. Improvements to the reconstruc-tion technique provided better resolution, but still left uncertainthe organization of individual heads [5,24,55,56], a situation thatwas only corrected by the use of cryo-EM together with a newmethod of reconstruction (see Section 4.3). The demonstration thatmyosin head helical order could be preserved in invertebrates wasfollowed by improvements in techniques for preserving helical or-der in vertebrate filaments as well, finally making possible 3Dreconstructions of these more complex structures [39,54,92].

4.3. Iterative Helical Real Space Reconstruction (IHRSR)

4.3.1. PrincipleHelical reconstructions are carried out in reciprocal space and

require computation of Fourier transforms, and identification andindexing of layer lines. This process is tedious, and determinationof helical symmetry can be problematic. An alternative, real-spacemethod – Iterative Helical Real Space Reconstruction (IHRSR) –also takes advantage of the multiple subunit views present in asingle image of a helix, but does so in a different way [93–95].An initial reference model of the structure is generated based on

construction of Limulus thin filament based on negative stain images. (a and b)Ao and Ai), and tropomyosin (Tm) associated with outer domain in low Ca2+ state (a)struction at low (c) and high (d) Ca2+. (a–d) Modified from [32], with permission. (e)ng ‘‘J’’-shaped structures in 4 co-axial helices represent pairs of myosin heads (see

Fig. 6. Fitting of myosin head atomic structure into repeating J-motif in tarantulareconstruction (Fig. 5e). Motor domain (MD), essential light chain (ELC) andregulatory light chain (RLC) of blocked head are in green, orange and yellow; thoseof free head are blue, pink and beige. Asterisk indicates rod-like volume of densitythat represents the start of the S2 portion of the myosin tail. Modified from [50].

R. Craig / Methods 56 (2012) 33–43 41

known information, for example a preliminary helical reconstruc-tion (Section 4.2). Density projections of the model are computedfor different rotations about its long axis (e.g. every 4�). Filamentimages are computationally boxed into short, overlapping seg-ments, and each segment thus produced is compared with the dif-ferent projections of the model by cross-correlation (typically thebox length is chosen to contain a small number of subunits ofthe repeating structure, and the box is shifted longitudinally byone subunit between segments: the optimal parameters are deter-mined by trial and error). The angle of the model that gives the bestmatch to the filament segment is then assigned to that segment. Areconstruction is computed from the different segments by backprojection using these assigned angles, and the reconstruction heli-cally averaged. The resulting reconstruction becomes the model fora second round of matching, and the process is repeated until thereare no further changes in the reconstruction. This iterative realspace technique has some key advantages over traditional helicalreconstruction. For example, filament straightening and layer linepicking are not required, and filaments with disorder or with par-ticular symmetries that are problematic for helical reconstruction(because information from different helical tracks overlaps in theFourier transform) can often be readily reconstructed by IHRSR.

4.3.2. UsesA major breakthrough in our understanding of thick filament

structure was obtained using IHRSR to reconstruct cryo-EM imagesof relaxed tarantula filaments [50]. The reconstruction had a reso-lution of 2.5 nm (a 2-fold improvement over the negative stainhelical reconstruction), revealing the myosin heads unambiguouslyand showing that the two heads of each myosin interacted witheach other (Fig. 5e; see Section 4.4). Subsequent studies of otherinvertebrate thick filaments and isolated myosin molecules haveshown that this interacting-head motif is likely to be widespreadacross the animal kingdom [50–53,96,97]. Comparable observa-tions on vertebrate filaments (mouse cardiac) have been muchmore difficult and are at lower resolution due to the more complexstructure and less stable myosin head arrays of vertebrates; how-ever, they too appear to show a similar structure [54]. Interpreta-tion of the complex vertebrate reconstruction depended stronglyon the insights first obtained from the unambiguous results ofthe invertebrate (tarantula) filaments. In addition to revealingthese head-head interactions, the tarantula reconstruction alsoshowed the subfragment 2 (S2) region of the myosin tail, and thepresence of subfilaments in the filament backbone. The assemblyof myosin tails into subfilaments had been predicted 25 years ear-lier on the basis of model building using invertebrate muscle X-raydiffraction data [98]; the tarantula reconstruction provided com-pelling support for this model. Currently the best thick filamentreconstruction has been limited to 2.5 nm resolution, due in partto the use of a conventional (non-field emission) microscope. Otherfilaments (e.g. F-actin) have been reconstructed to as high as0.6 nm resolution (enabling the visualization of secondary struc-ture such as a-helices) using field emission microscopy and moreadvanced image processing techniques [59,60]. Whether similartechniques applied to thick filaments are as successful will dependon whether inherent disorder in the heads caused by their flexibleattachment to the tails limits resolution.

4.4. Atomic fitting

A final and key step in the interpretation of reconstructions ofsufficient resolution is the computational fitting (docking) of atom-ic structures of filament subunits (e.g. myosin heads, actin mono-mers) into the reconstruction. If the fit is unambiguous, which ispossible when subunits are asymmetric, this effectively extendsthe resolution of the reconstruction to near-atomic level. Fitting

can be carried out using a number of (free) software packages, suchas UCSF Chimera [99] and SITUS [100]. The excellent fit of theinteracting-head structure present in crystals of smooth muscleheavy meromyosin [101] into the J-shaped motif (Fig. 5e) of thetarantula thick filament reconstruction directly demonstrated thepresence of similar interactions in the intact filament (Fig. 6)[50]. In this structure, one head (the ‘‘free’’ head) physically blocksthe actin-binding region of the other (the ‘‘blocked’’ head) – thushindering binding of the blocked head to actin – while the blockedhead is close to the converter domain and essential light chain ofthe free head, which could prevent the free head from functioningnormally during ATP turnover [50,101,102]. Thus a 2.5 nm resolu-tion reconstruction is interpreted in terms of specific polypeptideregions of the protein subunits, whose interaction suggests a sim-ple structural model for the switching off of myosin head activityin relaxed muscle. Similarly, the position of tropomyosin on thinfilaments in relation to specific actin residues was revealed by fit-ting of the atomic structure of actin into the 3D reconstruction ofLimulus thin filaments [32]. The results showed that in the lowCa2+ state, tropomyosin physically blocked clusters of amino acidresidues on actin known to be involved in myosin binding, whilethese residues became exposed when tropomyosin moved to itshigh Ca2+ position. These near-atomic level observations providedstrong support to the steric blocking model for thin filamentregulation.

Acknowledgments

I thank Dr. Raúl Padrón for comments on the manuscript. Theauthor’s work is supported by NIH Grants AR034711 andHL059408.

Appendix A

Preparing holey carbon films

1. Make up 100 ml of 0.5% (w/v) formvar in chloroform. Stirwith magnetic stirrer. Takes a while to dissolve (15–20 min).

2. Thoroughly mix 10 ml glycerol with 10 ml distilled water.

42 R. Craig / Methods 56 (2012) 33–43

3. Put 20 drops of glycerol/water mixture (from Pasteur pip-ette) into 50 ml of formvar solution. Use 50 ml stopperedgraduated cylinder.

4. Immediately prior to sonication, shake vigorously for 30 s.Shaking disperses the glycerol/water into fine droplets inthe chloroform/formvar, but the droplets are not smallenough and readily coalesce. The sonication step which fol-lows makes the droplets much smaller and the right size forthe holey film. This preliminary dispersion by shakingensures that the glycerol/water is not floating as a ‘‘glob’’at the surface when the sonication starts, in which case itis difficult to disperse.

5. Sonicate for 45–60 s at setting 6 using a microprobe sonica-tor (setting will depend on the sonicator and may be foundby trial and error). After sonication, the mixture should feelwarm and look uniformly milky (an emulsion of glycerol/water microdroplets in the formvar/chloroform).

6. Put solution into a staining (Coplin) jar. Clean several 3 � 1inch glass microscope slides with Ross (lint-free) lens paper,dip them one at a time into formvar solution for a couple ofseconds, remove, stand on their ends and leave until theyappear dry by eye. Do all of the slides at one time becausethe dispersion is unstable and the droplets slowly coalesce.If it is a very dry day and the holes are few or small, itmay help to dip and dry the slides in a humid chamber(about 60% RH).

7. As soon as one slide is dry, float the film off on water, pick upone test grid, dry it briefly (5 min), cut out the grid from theformvar with a razor blade, and check in the EM for hole size,number and distribution. For method, see (8) below. Thereshould be plenty (minimum of 10–20 per 400-mesh gridsquare) of variable sized holes, often patchily distributed.To test whether the holes are large enough, check in theelectron microscope at 50,000�: the larger holes should belarge enough so that their edges are outside the field of viewon the large fluorescent screen. The holes may have ‘‘ragged’’edges and may be crossed by fine, very thin strands of form-var. These will be removed with the methanol etching steplater. If the holes look good, proceed with the other slides,picking up 40–50 grids per slide (see (8,9) below). If theydo not look good, discard the slides and start again, varyingsonication time, humidity etc., until good holes are obtainedon a test grid.

8. Fill crystallizing dish with distilled water. Wipe surface ofwater clean by dragging a sheet of Ross lens tissue acrossit. With a clean razor blade, score the film on one face ofthe slide near the edges so that the film will easily detachin the next step. Carefully float off the film onto the watersurface by sliding the slide (scored side up) into the waterat a 45o angle. ‘‘Huffing’’ on the slide first with moist breathhelps to separate the film from the slide.

9. Place grids (no pre-cleaning necessary) face down on film onwater surface. Pick up with Whatman #4 filter paper or Bib-ulous paper by touching the paper horizontally to floatingformvar and then removing (these papers absorb water atthe right rate to facilitate pick-up of grids and film). Firstmake 90o bend at end of filter paper to act as handle.

10. After filter paper is dry, place it (with attached grids/form-var, grids uppermost) on a bed of filter paper saturated withmethanol in a glass Petri dish for 10 min. (Use fume hood).The bottoms of the grids should be soaked from underneath,but the grids should not be immersed in the methanol. Coverdish so that methanol does not evaporate. This ‘‘etching’’procedure removes glycerol/water droplets, dissolves anyvery thin plastic that might have been crossing ‘‘pseudo’’holes, and gives a sharper edge to the real holes. After the

grids are dry, check a grid in the EM to make sure that thishas occurred. In particular, check holes at high magnificationto make sure that they are real holes and not pseudo-holes(still having a thin coating of formvar stretching across thehole).

11. If the holes look good, coat the grids heavily with carbon (sothat the supporting filter paper appears dark brown).

12. Set up a large glass Petri dish in hood with a bed of filterpaper saturated with 1,2-dichloroethane (same procedureas for methanol) and place grids/filter paper/formvar/carbononto the wet filter paper in the Petri dish for 2 h (grid sideup, as in 10). This dissolves the formvar, leaving the holeycarbon. Grids without the formvar are more stable in theelectron beam (less drift, holes do not expand in beam, etc.).

13. Remove filter paper/grids and dry in a glass Petri dish. Storein a glass Petri dish in a refrigerator (glass dishes avoid elec-trostatic attraction of grids, which can be a problem withplastic dishes; in our experience, the grid surface remainscleaner and its surface properties remain more constantwith storage in a refrigerator).

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