Rhombomere Developmentin a Reptilian Embryo

MICHAEL B. PRITZ*Section of Neurological Surgery, Indiana University School of Medicine,

Indianapolis, Indiana 46202-5124

ABSTRACTRhombomere development was investigated in a reptile, Alligator mississippiensis, using

a variety of methodologies: cytoarchitecture (cresyl violet), histochemistry (peanut aggluti-nin), immunocytochemistry (antibodies to acetylated tubulin, vimentin, calretinin, andacetylcholinesterase), and external and internal morphology of wholemount embryos. Rhom-bomere boundaries form sequentially until 8 rhombomeres are present at stage 8. From stage11 onwards, rhombomere borders fade. When present, boundaries of rhombomeres 2 through5 were distinct. In all embryos, except the earliest stages, neural tissue was divided betweenthe caudal end of the mesencephalon and the rostral end of the rhombencephalon. This area oftransection was designated as the isthmus. For these technical reasons, a distinct borderbetween the midbrain and the first rhombomere was not seen and the isthmic rhombomerecould not be identified. The interrhombomeric boundary between rhombomere 7 andrhombomere 8 and between the most caudal rhombomere and the spinal cord was not nearlyas clear as were the boundaries of rhombomeres 2 through 5. Development of rhombomeres 2through 5 was investigated in wholemount preparations between stages 5/6 and 11.Qualitative and quantitative observations were made. In these rhombomeres, r2 through r5,rostrocaudal caudal expansion occurs at a slower rate than mediolateral development. Thisdifferential growth sculpts the morphology of rhombomeres 2 through 5.

Rhombomere development in Alligator shares several features in common with hindbrainsegmentation in chick. The identification of rhombomeres in a multitude of vertebrates from avariety of classes suggests that segmentation is a feature common to hindbrain developmentin all vertebrates. J. Comp. Neurol. 411:317–326, 1999. r 1999 Wiley-Liss, Inc.

Indexing terms: brainstem; embryology; evolution; hindbrain

How can alterations in embryonic development causeevolutionary change? Although such a question is not new(Roux, 1895, quoted in Gilbert et al., 1996), it has only beenrecently that technical and methodological advances haveallowed new approaches to address just such a problemfrom both a developmental as well as an evolutionaryperspective (Muller, 1991; Gilbert et al., 1996; Raff, 1996).

In the central nervous system, a developmental evolu-tionary approach to this type of problem has been exempli-fied by investigations into hindbrain development. Identi-fication of iterative units in the rhombencephalon,rhombomeres, was described over 100 years ago (Orr,1897). Since then, a multitude of descriptive and experi-mental studies have investigated hindbrain segmentationin a variety of vertebrates (for review, see Gilland andBaker, 1993). However, the bulk of these observations havefocused on representatives from two classes of vertebrates,chick and zebrafish (Gilland and Baker, 1993).

Although data from these two species (chick and ze-brafish) have provided considerable information regarding

rhombomeres and their development, these studies do notallow the decision as to whether similar observations inchick and zebrafish are due to independently evolvedfeatures or the result of inheritance from a commonancestor because birds and teleosts are only distantlyrelated.

In order to distinguish between these two possibilities,representative species from other vertebrate classes re-quire study. For this reason, the American alligator, Alliga-tor mississippiensis, was chosen as an experimental ani-mal because it is a member of the sole surviving reptiliangroup that shares a common ancestry with birds (Whet-

Grant sponsor: Indiana University Biomedical Research Fund and A. C.,B. M., and G. Fossa Spinal Cord Injury Research Fund.

*Correspondence to: Dr. M.B. Pritz, Section of Neurological Surgery,Indiana University School of Medicine, 545 Barnhill Drive, Emerson 139,Indianapolis, IN 46202-5124. E-mail: mpritz@iupui.edu

Received 28 July 1998; Revised 16 February 1999; Accepted 23 March1999

THE JOURNAL OF COMPARATIVE NEUROLOGY 411:317–326 (1999)

r 1999 WILEY-LISS, INC.

stone and Martin, 1979). Features common to both Alliga-tor and chick would suggest that these characters are dueto inheritance from a common ancestor rather than inde-pendently evolved features. Furthermore, charactersshared by these two amniotes (chick and Alligator) andzebrafish would imply that these traits are common tohindbrain development in all vertebrates rather thanindependent evolution on three separate occasions (North-cutt, 1990). If so, hindbrain organization in all vertebrateswould begin with a common plan that only later varied indevelopment to produce the different hindbrain morpholo-gies observed in species as diverse as frog and monkey.

The presence of rhombomeres in Alligator was firstnoted by Clarke (1891) and was subsequently documentedby others (Ferguson, 1985; Gilland and Baker, 1993).However, details as to the number of units, sequence ofboundary formation, and other descriptive features ofhindbrain development are lacking. The present analysisexamines this problem using a variety of descriptive(Vaage, 1969) and experimental approaches that havesuccessfully identified rhombomeres in other vertebrates(Heyman et al., 1995; Layer and Alber, 1990) as well asothers that have not been specifically used. A preliminaryaccount of some of these findings has been presentedpreviously (Pritz, 1996).

MATERIALS AND METHODS

The protocols described below were approved by theanimal care committee at Indiana University School ofMedicine . These details conformed to National Institute ofHealth guidelines.

Animals

Eggs were obtained from the Rockefeller Wildlife Refugein Grand Chenier, Louisiana as a clutch containing 38–53eggs all laid simultaneously. Before incubation, all eggswere candled and the location of embryos marked. Viableembryos were incubated at 30°C in a 2:1 mixture ofvermiculite and water with the embryo positioned on top.Under these conditions, embryos will hatch in 62–67 days(Ferguson, 1985).

Embryos were killed between stages 2 and 15. First,eggs were candled to identify the embryo location. Then,the eggshell surrounding the embryo was opened with finescissors under an operating microscope. Identified, viableembryos were placed in fixative. A variety of fixatives wereused for embryos processed for histochemistry or immuno-cytochemistry. Best results were obtained by placement in100% methanol where embryos could be stored at 280° Cfor many months. Prior to prolonged storage, embryoswere staged according to the scheme of Ferguson (1985).Except at the earliest stages, brains were dissected freefrom the surrounding tissue including cranial nerves andotic vesicle/otocyst using fine tungsten needles, jeweler’sforceps, and microscissors, under a dissecting microscope.Except at earliest stages, brains were divided into twopieces at the isthmus, the constriction between the caudalmesencephalon and rostral rhombencephalon (Lumsdenand Krumlauf, 1996). One contained the hindbrain,whereas the other consisted of the midbrain and forebrain.The roof plate and extraneural membranes were divided soas to allow the hindbrain to lay flat. In brains stored at

280°C, the best histochemical and immunocytochemicalresults were obtained by gradual rewarming of tissue:220°C (30–60 minutes); 4°C (30–60 minutes); and thenroom temperature before embedding. Brains processed forcresyl violet staining were placed in a variety of fixatives:Bouin’s solution; 10% foramalin; 4% paraformaldehyde; or4% glutaraldehyde.

Embedding and sectioning

Embryos were embedded in gelatin or albumin-gelatin.Blocks processed for histochemical or immunocytochemi-cal methodologies were placed in a solution of 30% sucrose-sodium phosphate buffer (PBS; 0.1 M at pH 7.2) overnightat 4°C. To prevent the embedding medium from dissolvingduring tissue processing, blocks were placed in a solutionof 2% formalin-sucrose-PBS for 4 hours prior to sectioning.Blocks of brains stained for cresyl violet were placed in asolution of 30% sucrose-formalin or 30% sucrose-4% glutar-aldehyde overnight before sectioning.

Frozen sections were cut longitudinally or sagittally at25 or 30 µm on a sliding microtome and collected in PBS.Transverse sections were not useful at the stages ofdevelopment examined.

Peanut agglutinin histochemistry

Free-floating sections were washed in three changes ofPBS, incubated for 12–24 hours in horseradish peroxidase(HRP)-peanut lectin (50 mg/ml), washed in three changesof PBS, and then incubated in the following diaminobenzi-dine (DAB) solution: 100 mg DAB in 10 ml of distilledwater plus 1 ml of Triton X-100 plus 20 ml of 0.2 Mcacodylic buffer (pH 5.6) plus 6 µl of 30% hydrogenperoxide for 30 seconds. The reaction was stopped bytransferring the sections to 70% alcohol for 1 minute.Sections were then stored in PBS at 4°C until they weremounted out of distilled water onto chrome alum-coatedslides and air-dried. Mounted sections were then dehy-drated through a series of graded alcohols, cleared inxylene, and coverslipped.

Immunocytochemistry

Successful results were obtained with the followingantibodies: acetylated tubulin, vimentin, calretinin, andacetylcholinesterase. Methodology was similar except forprimary antibody concentration, secondary antibody, andperoxidase-antiperoxidase complex. In the case of primaryantibodies raised in mouse (acetylated tubulin, vimentin,and acetylcholinesterase), free-floating sections werewashed in three changes of PBS and then incubated inprimary antibody in PBS containing 2% normal goatserum. The dilutions of the primary antibodies were:monoclonal anti-acetylated tubulin (Sigma, St. Louis, MO),1:500; monoclonal anti-vimentin (Sigma), 1:100 or 1:50;and monoclonal anti-acetylcholinesterase (Chemicon, Tem-ecula, CA), 1:100. Then, sections were transferred throughthree changes of PBS and incubated with secondaryantibody (goat anti-mouse IgG, Vector Labs, Burlingame,CA) in PBS containing 2% normal goat serum at aconcentration of 1:100 for 1 hour followed by three changesof PBS. Sections were then incubated in mouse peroxidase-antiperoxidase complex (Chemicon International) in PBScontaining 2% normal goat serum at a dilution of 1:300 for1 hour. Sections were then washed in three changes of PBS

318 M.B. PRITZ

and reacted for 30 seconds in a DAB solution consisting of:100 mg DAB in 10 ml of distilled water plus 1 ml of TritonX-100 plus 20 ml of 0.2 M cacodylic buffer (pH 5.6) plus 6 µlof 30% hydrogen peroxide. The reaction was stopped bytransfer to a solution of 70% alcohol for 1 minute. Sectionswere stored in PBS at 4°C until they were mounted out ofdistilled water onto chrome alum-coated slides and thenair-dried. Mounted sections were then dehydrated througha series of graded alcohols, cleared in xylene, and thencoverslipped.

Calretinin immunocytochemistry methodology was iden-tical to that described above for antibodies raised in mouseexcept for the following changes. The primary antibodywas rabbit polyclonal anti-calretinin (Chemicon) at aconcentration of 1:500; the secondary antibody was goatanti-rabbit IgG (Vector Labs); and the tertiary antibodywas rabbit peroxidase-antiperoxidase complex (Chemi-con).

The following antibodies failed to label rhombomeres:monoclonal mouse anti-neural cell adhesion molecule (N-CAM; Sigma); monoclonal mouse anti-growth-associatedprotein-43 (GAP-43; Sigma); and monoclonal mouse anti-glial fibrillary acid protein (GFAP; Boehringer Mannheim,Indianapolis, IN). Controls for nonspecific staining substi-tuted normal mouse or rabbit serum for the primaryantibody at a concentration equal to or greater than that ofthe primary antibody. Rhombomeres were not visualizedin tissue processed in this fashion.

Wholemount preparations

After hindbrains had been freed of surrounding tissueby using a dissecting microscope, they were routinelyexamined for the presence of rhombomeres. Selected hind-brains were then photographed by using indirect lightthrough a Zeiss microscope as viewed from either theexternal or internal (ventricular) surface.

In order to examine rhombomere development overtime, wholemount hindbrains stored in 4% glutaraldehydefrom a single clutch were examined at a time whenrhombomeres 2 through 5 were clearly visible. Hindbrainwholemounts were studied and drawn from the perspec-tive of the internal surface (ependymal side) at 43 magni-fication using an Olympus BH-2 microscope. A cameralucida attachment was used to draw outlines of representa-tive hindbrain wholemounts for qualitative observations.

For quantitative analyses, outlines of rhombomeres 2through 5 were drawn. Measurements of these rhombo-mere profiles were made by using an image analysissystem (NIH image l.61) attached to a personal computer.For each embryo, two sets of rhombomere profiles wereobtained. The interrhombomeric border was not includedin rhombomere profile measurements. The following mea-surements were made on rhombomere profiles: area, length(measurement of longest axis), width (measurement per-pendicular to length), and ‘‘eccentricity’’ which was a ratioof width/length.

To provide some quantitative values for the qualitativeobservations of rhombomere development, the areas ofrhombomere profiles 2 through 5 illustrated in Figure 2were summed and the percentage for each given rhombo-mere area profile for this series of embryos was calculated.Throughout the text and tables, measurements were ex-pressed as mean 6 standard error (S.E.). No correction fortissue shrinkage was made.

Histological analysis

Sectioned tissue was viewed using standard light micros-copy using brightfield illumination or Nomarski optics.Appropriate sections were drawn using a camera lucidaattachment or photographed.

RESULTS

A variety of approaches were used to identify rhombo-meres, interrhombomeric borders, as well as rhombomeredevelopment and disappearance. Rhombomeres 2 (r2)through 6 (r6) are easily recognized (Gilland and Baker,1993) both in unstained wholemount preparations (Fig. 1)as well as in tissue sectioned longitudinally (Figs. 3 and 4)by using a variety of methods detailed below. The neuroepi-thelial tissue rostral to r2 and caudal to the isthmus(midbrain-rhombencephalon border) was designated asthe first rhombomere (r1). No attempt was made tosubdivide r1 further. Rhombomeres caudal to the sixthrhombomere and rostral to the spinal cord were desig-nated as rhombomeres 7 (r7) and 8 (r8).

Wholemount embryos

Transverse segments were clearly visible as early asstage 4. Interrhombomeric boundaries were recognized insome embryos through stage 11. Subsequently, bordersbetween rhombomeres began to fade. Rhombomeres were

Fig. l. Wholemount photographs of unstained hindbrains transec-ted at the isthmus from a stage 8/9 Alligator embryo viewed frominternal (A) and external (B) surfaces. Orientation arrows are similarfor each view. Arrowheads mark interrhombomeric divisions thatidentify rhombomeres 2 (r2) through 6 (r6). c, caudal; m, medial. Scalebar 5 500 µm.

HINDBRAIN DEVELOPMENT IN ALLIGATOR 319

not evident at stage 14/15. Rhombomeres 2 through 5 weredistinct beginning at stage 5/6 (Fig. 2) and could be seeneither from either the internal (ventricular; Fig. 1A) or theexternal surface (Fig. 1B). Although rhombomere 6 couldbe seen at early stages (6/7 through 9/10; Fig. 2), thisboundary could not be recognized at stage 10 and beyond.Clear identification of r7 and r8 was never seen regardlessof the surface examined. Similarly, r1 was identified onlyas the neuroepithelial tissue caudal to the isthmus butrostral to r2.

To provide a perspective of rhombomere growth overtime, outlines of rhombomeres 2 through 5, which remaindistinct throughout rhombomere development, were drawn(Fig. 2). The progressive change in shape and size ofrhombomeres 2 through 5 and their interrhombomericborders over time is clearly evident (Fig. 2). To providedata as to the relative proportion of an individual rhombo-mere area in relation to the entire (summed) area ofrhombomeres 2 through 5 over time, measurements ofareal profiles of individual rhombomeres were made (Table1) for the example illustrated in Figure 2. Initially, r2 is thelargest rhombomere, whereas r5 is the smallest with r3and r4 being nearly equal. As time passes, r3 increases insize, whereas r4 becomes proportionally the smallestrhombomere with r5 remaining at approximately the sameproportion at the oldest stage of development (stage 11).Rhombomere 2 slightly decreases its proportional area atstage 11 as compared with stage 5/6.

To provide quantitation for the growth, development,and shape of rhombomeres 2 through 5 over this sameinterval, stages 5/6 through 11, measurements of area(Table 2), and eccentricity (Table 3) were made on drawnprofiles. Although these profile measurements of area(Table 2) parallel the quantitative (Table 1) data of themeasurements made on the developmental sequence illus-trated in Figure 2, some differences occurred (Table 2)because r2 through 5 were drawn from several embryosand each embryo provided two sets of data. Nevertheless,these quantitative observations indicate tremendousgrowth in r2 through 5. This has differentially affectedthese rhombomeres. Areal profiles have increased as fol-lows: 3.5 times for r3, slightly more than 3 times for r2; andslightly less than 3 times for r5; whereas the final growth

Fig. 2. Camera lucida drawings of wholemount, right side hindbrains of Alligator embryos drawn fromthe internal surface. The development of rhombomeres 2 through 5 are illustrated from stages 5/6through 11. c, caudal; m, medial; sc, spinal cord; 2–5, rhombomeres 2–5. Scale bar 5 500 µm for allhindbrain sections.

TABLE 1. Relative Area Sizes of Individual Rhombomeres1 2 Through 5During Development

Stage r2 (%) r3 (%) r4 (%) r5 (%)

5/6 29.4 24.3 24.0 22.36/7 30.3 25.5 21.5 22.77/8 30.1 26.1 19.9 23.98 26.6 28.5 20.5 24.49 29.9 25.7 20.3 24.1

9/10 27.3 28.6 19.1 25.010 27.3 33.1 16.1 23.511 26.9 31.6 19.0 22.5

1r, rhombomere.

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of r4 is only about 2.4 times its initial areal profile value(Table 2). Furthermore, these areal profile changes fromstage to stage can be seen.

The eccentricity data (Table 3) from drawn profilesprovides a quantitative way to view the shape of individualrhombomeres 2 through 5 illustrated in Figure 2. Aneccentricity value of 0.5 describes a rectangle whose lengthis twice that of its width. As this value approaches 1, theshape of a rhombomere profile approximates a square. Onthe other hand, as eccentricity values reach 0, rectangularshapes become more eccentric. Initially, whereas eachrhombomere is rectangular in shape, the variation be-tween the width (rostrocaudal extent) and length (medio-lateral extent) are not substantially different at stage 5/6(Fig. 2; Table 3). However, as time passes, profile widthsand lengths grow at different rates so that by stage 11 (Fig.2; Table 3), except for r3, rhombomere profile length isnearly twice that of rhombomere width. For rhombomeresr2 through r5, profile shape becomes more eccentric overtime. This implies that rostrocaudal growth occurs at arate slower than mediolateral growth.

Rhombomere identification basedon staining patterns

Although tissue processed in the sagittal plane dididentify rhombomeres, hindbrain segmentation was mostclearly visible in tissue sectioned longitudinally. Accord-ingly, the vast majority of hindbrains were examined fromthis perspective. Nissl staining (Fig. 3A–D), peanut agglu-tinin histochemistry (Fig. 3F) and immunocytochemistryusing an antibody to acetylated tubulin (Fig. 3E) proved tobe the most distinct markers for hindbrain segment identi-fication. Antibodies to calretinin (Fig. 4A), vimentin (Fig.4B), and acetylcholinesterase (Fig. 4C) also clearly re-vealed rhombomeres. Identification of rhombomeres wasbetter visualized with calretinin than with vimentin oracetylcholinesterase. However, tissue processed for Nisslstain, peanut agglutinin histochemistry, and acetylated

tubulin immunocytochemistry gave the best results. Thus,most of the observations were based on these lattermethodologies. Good correlation between observationsbased on these different methodologies (Nissl staining,peanut agglutinin histochemistry, and immunocytochemis-try with antibodies to acetylated tubulin, vimentin, calreti-nin, and acetylcholinesterase) was seen.

Rhombomere development

Except at earliest stages, tissue and cranial nervessurrounding hindbrains were dissected free from speci-mens processed for histological analysis. Thus, observa-tions noting the relationship between individual rhombo-meres and specific cranial nerves were not made nor couldthe relationship between otocyst location and a specificrhombomere be done. Determination of the developmentalsequence of hindbrain segmentation was undertaken bybeginning analysis at stage 6 where rhombomeres 2through 6 were distinct. Further investigation was thenundertaken by examining earlier and later stages. Interpre-tation at stages later than stage 6 (Fig. 3D–F) wasstraightforward, whereas analysis at earlier times (Fig.3A–C) was not. Although interrhombomeric boundarieswere present at stages earlier than stage 6, identificationof specific rhombomeres based solely on morphologicalcriteria required interpretation (Fig. 3A–C). When rhombo-meres 1 through 8 were not distinct, rhombomeres werecombined and marked accordingly (Fig. 3B–D). For ex-ample, when no distinction between hindbrain segmentswas present, they were marked as either r7 & r8 (Fig. 3D)or r6–8 (Fig. 3B and C).

Hindbrains as early as stage 1 were not available foranalysis. Reliable segment identification could be made asearly as stage 2 (Fig. 3A). At this stage, three subdivisionswere identified. These were designated as a, b, and c andtheir relationship to the otic pit can be seen. At stage 3(Fig. 3B), interrhombomeric borders were visible and fiveunits were identified. The first rhombomere with distinctrostral and caudal boundaries was interpreted as r2 andthe next most caudal segment as r3. Segments r4 and 5were interpreted as being fused, r4–5, as were segmentsr6–8. The first rhombomere, r1, was identified as neuraltissue rostral to r2. The caudal border of r1 was distinct,whereas the rostral boundary, for technical reasons (be-cause the hindbrain was divided at the isthmus), was notseen. At stage 4 (Fig. 3C), 6 rhombomeres were identified.At this time, r4 and r5 were interpreted as being subdi-vided and distinct, whereas boundaries between rhombo-meres 6 through 8 were not observed. By stage 6, 7rhombomeres were seen. Rhombomere 6 was distinct,whereas no border separating r7 and r8 was observed.Differentiation between rhombomeres 7 and 8 was seen asearly as stage 8 and became clear at stage 8/9 (Fig. 3E andF). By stage 11, rhombomere boundaries began to fade,and by stage 14, hindbrain segments could no longer bevisualized. The development and disappearance of rhombo-meres is illustrated schematically (Fig. 5).

Interrhombomeric borders

The interrhombomeric borders of r2 through r6 wereclear (Fig. 3). Because all but the earliest embryos weredivided at the isthmus, the constriction between thecaudal mesencephalon and the rostral rhombencephalon(Lumsden and Krumlauf, 1996), a distinct boundary be-tween the mesencephalon and r1 was not observed be-

TABLE 2. Mean Area Profile Measurements of Rhombomeres 2 Through 5Between Stages 5/6 and 111

Stage N

Mean area profiles (mm2 6 S.E.)

r2 r3 r4 r5

5/6 2 148.72 6 .08 120.87 6 2.61 112.90 6 9.03 109.56 6 3.406/7 2 186.76 6 5.99 166.31 6 4.60 136.36 6 3.23 142.11 6 2.247/8 4 195.18 6 6.18 179.04 6 7.59 139.81 6 3.05 159.69 6 5.058 6 233.78 6 3.17 235.10 6 5.51 175.88 6 4.85 202.51 6 3.139 4 268.38 6 14.83 264.49 6 4.84 203.51 6 4.58 237.03 6 3.28

9/10 4 312.43 6 5.88 352.27 6 5.07 246.59 6 6.43 288.06 6 3.1910 4 344.25 6 16.15 400.16 6 19.72 235.31 6 10.28 284.95 6 8.1411 2 366.66 6 1.56 421.91 6 11.34 269.59 6 9.10 309.17 6 .49

1N, number of rhombomere area profiles measured; r, rhombomere; S.E., standard error.

TABLE 3. Mean Eccentricity Profile Measurements of Rhombomeres 2Through 5 Between Stages 5/6 and 111

Stage N

Mean eccentricity profiles (6 S.E.)

r2 r3 r4 r5

5/6 2 0.75 6 .003 0.75 6 .002 0.68 6 .015 0.79 6 .0296/7 2 0.66 6 .008 0.69 6 .019 0.67 6 .029 0.76 6 .0297/8 4 0.64 6 .005 0.73 6 .018 0.69 6 .023 0.76 6 .0308 6 0.60 6 .006 0.69 6 .008 0.62 6 .010 0.66 6 .0109 4 0.63 6 .020 0.67 6 .019 0.62 6 .011 0.67 6 .020

9/10 4 0.52 6 .024 0.63 6 .002 0.54 6 .018 0.59 6 .01210 4 0.52 6 .025 0.62 6 .018 0.49 6 .025 0.55 6 .00511 2 0.53 6 .008 0.61 6 .030 0.49 6 .013 0.54 6 .004

1N, number of rhombomere profiles measured; r, rhombomere; S.E., standard error.

HINDBRAIN DEVELOPMENT IN ALLIGATOR 321

cause tissue was transected at this level. Furthermore, theinterrhombomeric borders of r7 and r8 (Fig. 3E and F),when present, were not nearly as clear as were thosebetween the borders of r2 through r6. In addition, tissuestained with acetylated tubulin (Fig. 3E) more clearlyvisualized rhombomere borders than did peanut aggluti-nin (Fig. 3F), calbindin (Fig. 4A), vimentin (Fig. 4B), oracetylcholinesterase (Fig. 4C). A sharp demarcation be-tween the most caudal rhombomere and the spinal cordwas not commonly seen (Fig. 3).

Rhombomere boundaries visualized by peanut aggluti-nin histochemistry (Figs. 3F and 6A) were formed byfibers. These boundaries were visible in cresyl violet-stained sections (Fig. 3A–D), and were immunoreactive foracetylated tubulin (Figs. 3E and 6C), vimentin (Figs. 4Band 6B), acetylcholinesterase (Fig. 4C), and calretinin(Fig. 4A). In immunocytochemical material using antibod-

ies to acetylated tubulin, immunoreactive fibers at moredorsal levels (closer to the internal or ependymal surface)were oriented medial to lateral (Fig. 6C, right) in tissuesectioned longitudinally and then fanned out rostrocaudalat deeper levels to become fiber-free (Fig. 6C, left, and Fig.3E).

DISCUSSION

These morphologic observations have identified the num-ber, sequence of development, interrhombomeric borders,and morphology of iterative units in the hindbrain ofAlligator embryos using a variety of techniques betweenstages 2 and 15. Interpretation of these findings and therelation to hindbrain segmentation in other vertebratesare discussed further.

Fig. 3. Morphology of rhombomere development. Longitudinalsections through the right hindbrain of Alligator embryos at stages 2(A), 3 (B), 4/5 (C), 7 (D), 8/9 (E), and 10 (F) stained with cresyl violet(A–D), an antibody to acetylated tubulin (E), and by peanut agglutininhistochemistry (F). Arrowheads (A–F) indicate distinct interrhombo-meric borders, whereas small arrows (B, E, F) mark less obvious

boundaries between rhombomeres. Large arrows (D) denote an inter-rhombomeric boundary that does not extend through the entiremediolateral rhombomere width. 1–8, rhombomeres 1–8; a–c, rhombo-meres a–c; l, lateral; m, mesencephalon; op, otic pit; p, prosencephalon;r, rostral; sc, spinal cord. Orientation for all sections (A–F) is identical.Scale bars 5 200 µm for A–F.

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Developmental significance of rhombomeres

For neuromeres in general, and rhombomeres in particu-lar, to be of developmental significance at least some of thefollowing criteria have been suggested as requirements(Keynes and Lumsden, 1990; Lumsden, 1991). First, theneuromeric pattern should correspond to a segmentalpattern of molecular and/or cellular differentiation. Sec-ond, the neuromeric pattern should be produced by changesin cell proliferation and/or cell shape. Third, bordersbetween segments should create lineage restrictions andthus represent barriers to cell movement. Fourth, geneswith regulatory roles during development should be ex-pressed in relation to the neuromeric pattern.

Although segmentation in the developing hindbrain hasbeen documented in representatives of a wide variety ofvertebrates (Gilland and Baker, 1993), only in chick andzebrafish have these problems been extensively studiedwith modern techniques. Rhombomeres begin as a two-segment unit that later subdivides sequentially. Thisresults in eight segments in chick (Vaage, 1969), whereasnine have been identified in zebrafish (Hanneman et al.,1988). The first seven of these segments in zebrafish areclear, whereas the caudal units are less distinct (Hanne-man et al., 1988). The rostral seven rhombomeres in chickand zebrafish are felt to be homologous (Keynes andLumsden, 1990), whereas the posterior segments in ze-brafish were referred to as r8 of chick (Keynes andLumsden, 1990).

The notion of a two-segment periodicity was initiallybased upon the exits of cranial nerve location in relation toindividual rhombomeres and the correspondence betweenthe rhombomere location of branchial motor cell bodiesand individual branchial arch location (Lumsden andKeynes, 1989; Kuratani and Eichele, 1993). This periodic-ity has resulted in an alternate pattern of features thatdistinguishes odd- from even-numbered rhombomeres.Such characters include: appearance of neurons and axonswithin rhombomeres (Lumsden and Keynes, 1989), neuro-filament staining (Lumsden and Keynes, 1989), HNK-1antibody immunoreactivity (Kuratani and Eichele, 1993),and differential cell affinity (Wizenmann and Lumsden,1997). Further support for a segment periodicity has comefrom transplantation experiments (Guthrie and Lumsden,1991; Guthrie et al., 1993) and mutational analysis (Moenset al., 1996). Another feature of rhombomeres is that theirboundaries represent cell lineage compartments (Fraser etal., 1990). However, these boundaries are not impen-etrable (Birgbauer and Fraser, 1994). Lastly, expression ofcertain regulatory genes is correlated with rhombomericposition (for review, see Lumsden and Krumlauf, 1996;Prince, 1998).

When compared with these considerable data, the pre-sent observations have been limited to certain features ofrhombomere formation and development in Alligator.Rhombomeres and their borders were identified in cresyl

Fig. 4. Photograph of longitudinal sections of the left hindbrain ofAlligator embryos illustrating rhombomeres and interrhombomericboundaries stained with antibodies to calretinin (A; stage 9/10 em-bryo), vimentin (B; stage 10 embryo), and acetylcholinesterase (C;stage 7 embryo) viewed with brightfield (A) and Nomarski (B and C)optics. Interrhombomeric boundaries are identified laterally (arrow-heads) and medially (arrows). c, caudal; m, medial. Orientation foreach section is similar. Scale bars 5 100 µm for A–C.

HINDBRAIN DEVELOPMENT IN ALLIGATOR 323

violet-stained material. Rhombomere boundaries were vi-sualized by markers that labeled fibers (antibodies toacetylated tubulin and vimentin) as well as by peanutlectin-binding glycoprotein. Similar features have alsoidentified interrhombomeric borders in chick (Guthrie,1996; Heyman et al., 1995; Layer and Alber, 1990) and inzebrafish (Trevarrow et al., 1990). Other techniques thatalso marked rhombomeric boundaries in Alligator wereantibodies to calretinin and acetylcholinesterase whichconfirmed the observations made with the techniquesdescribed above, but which have not been extensively usedin other vertebrates.

These methods formed the basis to analyze Alligatorembryo hindbrains between Ferguson (1985) stages 2 and15. At stage 6, rhombomeres 2 through 6 were clear.Rhombomere 1 was designated as that segment rostral tor2 and caudal to the isthmus. Because tissue was dividedat the isthmus, the midbrain-hindbrain junction, therostral border of r1 was not observed and the identificationof an isthmic segment, rhombomere 0, noted in chick(Vaage, 1969), was not possible. Neural tissue between thecaudal end of r6 and the spinal cord was labeled asrepresenting r7 and r8. By stage 8, a separation betweenr7 and r8 was identified; however, this border was notnearly as evident as other interrhombomeric boundaries.Similarly, the hindbrain-spinal cord junction was not wellvisualized by any technique. Determination of rhombo-meres at stages earlier than stage 6 is open to interpreta-tion. The earliest stage available for analysis was stage 2where three segments were noted. These were merelylabled as a, b, and c, because their relationship to theultimate formation of rhombomeres was uncertain. Theseearly segmental units in Alligator may be similar toprorhombomeres described in chick (Vaage, 1969) andmouse (Osumi-Yamashito et al., 1996) because segment bin Alligator and prorhombomere b in chick (Vaage, 1969)and mouse (Osumi-Yamashita, 1996) all lie opposite theotic placode. How these segments in Alligator relate toprotosegements proposed in zebrafish (Moens et al., 1996)

is uncertain. Furthermore, it is likely that an early devel-opmental stage in Alligator exists where only two hind-brain units are present, as well as an even earlier stagetime when segmentation is absent.

The pattern of rhombomere formation between stages 2and 6 is open to question. Because tissue surrounding thehindbrain was removed, the location of the otic placode,cranial nerves, and cranial ganglion as well as the relation-ship between cranial nerves and ganglia and branchialarches could not be ascertained. However, this informationalone, even if available, may not have been sufficient tosolve this problem because certain cranial nerve motorneu-ron associations with rhombomeres varies among verte-brates (Gilland and Baker, 1993). Although some of thisvariation may result from cell migration as is the case forcertain branchiomotor nuclei in zebrafish (Chandrasekharet al., 1997), other techniques to identify rhombomeres willlikely be required to confirm individual rhombomere iden-tity. Nevertheless, and assuming that the early stages ofAlligator development are correct, differences betweenrhombomere formation between Alligator and chick wouldbe present. Assuming a one-to-one homology of individualrhombomeres between Alligator and chick, rostral rhombo-mere (rl–3) segmentation occurs earlier in hindbrain devel-opment in Alligator than it does in chick (Vaage, 1969). Iftrue, this may be reflected in a structural change in therostral hindbrain between Alligator and chick which maybe correlated with their respective ecological niches andbehaviors.

Rhombomere growth and development

Wholemount observations on the development of rhom-bomeres 2 through 5 document both qualitative changes inshape and quantitative alterations in a variety of mea-sured parameters over time. Similar studies in othervertebrates are limited. However, a recent report on chickhindbrain development (Kulesa and Fraser, 1998) is rel-evant to the present study on Alligator. These findings in

Fig. 5. Hindbrain development. Sequence of selected features inAlligator hindbrain development is shown diagramatically in longitu-dinal sections in which each stage has been drawn to the same lengthand proportions. Segmentation (labeled as a, b, and c) is observed atthe earliest stages available (stage 2). At stages 3 through 11,

rhombomeres 1 (rl) through 8 (r8) are seen. Clear separation andidentification of rhombomeres 2 (r2) through 6 (r6) are seen at stage 6.Boundaries between rhombomeres 7 (r7) and 8 (r8) are visualized atstage 8. Interrhombomeric borders begin to disappear at stage 11. Bystage 14, rhombomere boundaries have disappeared.

324 M.B. PRITZ

chick were, for the most part, made by time-lapse videomicroscopy on whole embryo explants, and used criteriadifferent from the present report to measure rhombo-meres. Nevertheless, data on chick hindbrain segmenta-tion (Kulesa and Fraser, 1998) indicate that rostrocaudallength of rhombomeres remains constant, whereas shapechanges are influenced by lateral expansions of the neuro-epithelium. Similarly, in Alligator, mediolateral expansionof r2 through 5 occurs more rapidly than does rostrocaudalgrowth.

Hindbrain segmentation in Alligator andother reptiles

Brief reference to hindbrain segmentation in Alligatorwas first made over 100 years ago (Clarke, 1891). Subse-quent observations documenting rhombomeres have beennoted in wholemount preparations (Ferguson, 1985) andby scanning electron microscopic techniques (Gilland andBaker, 1993). However, details of rhombomeric develop-ment and organization are limited.

Hindbrain segmentation has been investigated in rep-tiles other than Alligator. These include studies in lizards(McClure, 1890; Orr, 1887), snakes (Herrick, 1892), andSphenodon (Wyeth, 1924). The most detailed descriptionshave been made in Anolis where five (Orr, 1887) and six(McClure, 1890) rhombomeres have been identified. Fur-thermore, the relationship between rhombomeres andcranial nerves have been described (McClure, 1890; Orr,1887). Nevertheless, considerable details are lacking, mak-ing comparisons difficult between individual rhombomeresidentified in the present study and those identified byothers (McClure, 1890; Orr, 1887).

Rhombomeres in other vertebrates

The present observations have documented rhombo-mere development in Alligator. These findings support theobservations of others (Clarke, 1891; Ferguson, 1985;Gilland and Baker, 1993) that hindbrain segmentation ispresent in Alligator. The identification of rhombomeres ina multitude of vertebrates from a variety of differentclasses (Gilland and Baker, 1993) suggests that rhombo-meres are features common to the developing hindbrain ofall vertebrates. However, it remains to be seen whether thedevelopmental mechanisms to produce these iterativeunits in the hindbrain are similar among vertebrates andhow individual rhombomeres in various vertebrates com-pare. A knowledge of how these differences and similaritiesin rhombencephalon development have occurred mightenable the understanding of the variation in hindbrainmorphology observed in adult vertebrates that has beenproduced by evolution.

Fig. 6. Rhombomere boundary morphology. Camera lucida draw-ings of similarly oriented longitudinal sections of Alligator embryosstained by peanut agglutinin histochemistry (A; stage 6 embryo); withan antibody to vimentin (B; stage 10 embryo); and with an antibody toacetylated tubulin (C; stage 6 embryo) visualize interrhombomericboundaries. In C, because of section obliquity, acetylated tubulin-immunoreactive fibers are oriented parallel to the rhombomere bound-ary (right) and then fan out rostrally and caudally as the interrhombo-meric border becomes fiber free (left). Interrhombomeric boundariesare marked laterally (large arrowheads) and medially (small arrow-heads). c, caudal; m, medial. Scale bar 5 200 µm.

HINDBRAIN DEVELOPMENT IN ALLIGATOR 325

ACKNOWLEDGMENTS

I thank Dr. R. Elsey and J. Manning for generous supplyand donation of Alligator eggs; Drs. V. Lance, J. Lang, andM. Ewert for suggestions in raising and harvesting em-bryos; K. Si for technical assistance; D. Jaynes, J. Murphy,and Dr. L.C. Triarhou for help with photography; M.Houser and L. Green for manuscript preparation, and M.Williams for assistance with Figure 5. I am particularlygrateful to R.G. Northcutt for advice and encouragement,R. Baker for helpful suggestions on an earlier draft of thismanuscript, and two anonymous reviewers for their com-ments.

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