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Developmental Biology
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Proliferation and recapitulation of developmental patterning associated withregulative regeneration of the spinal cord neural tube
Gabor Halasi a,b, Anne Mette Søviknes a,1, Olafur Sigurjonsson a,2,3, Joel C. Glover a,b,⁎a Laboratory of Neural Development and Optical Recording (NDEVOR), Department of Physiology, Institute of Basic Medical Sciences, Domus Medica, University of Oslo, PB 1103 Blindern,0317 Oslo, Norwayb Norwegian Center for Stem Cell Research, Department of Immunology and Transfusion Medicine, Division of Diagnostics and Intervention, Oslo University Hospital-National Hospital,Domus Medica, PB 1112 Blindern, 0317 Oslo, Norway
⁎ Corresponding author at: Laboratory of Neural Deve(NDEVOR), Department of Physiology, Institute of Basicica, University of Oslo, PB 1103 Blindern, 0317 Oslo, No
E-mail address: [email protected] (J.C. Glov1 Present address: University of Bergen, Departme
N-5020 Bergen, Norway.2 Present address: The Blood Bank, Landspitalinn, S
Iceland.3 Present address: School of Science and Engin
Developmental patterning during regulative regeneration of the chicken embryo spinal neural tube was char-acterized by assessing proliferation and the expression of transcription factors specific to neural progenitorand postmitotic neuron populations. One to several segments of the thoracolumbar neural tube were selec-tively excised unilaterally to initiate regeneration. The capacity for regeneration depended on the stage whenablation was performed and the extent of tissue removed. 20% of surviving embryos exhibited complete reg-ulative regeneration, wherein the missing hemi-neural tube was reconstituted to normal size and morphol-ogy. Fate-mapping of proliferative adjacent tissue indicated contributions from the opposite side of theneural tube and potentially from the ipsilateral neural tube rostral and caudal to the lesion. Application ofthe thymidine analog EdU (5-ethynyl-2′-deoxyuridine) demonstrated a moderate increase in cell prolifera-tion in lesioned relative to control embryos, and quantitative PCR demonstrated a parallel moderate increasein transcription of proliferation-related genes. Mathematical calculation showed that such modest increasesare sufficient to account for the amount of regenerated tissue. Within the regenerated neural tube the expres-sion pattern of progenitor-specific transcription factors was recapitulated in the separate advancing ventraland dorsal fronts of regeneration, with no evidence of abnormal mixing of progenitor subpopulations, indi-cating that graded patterning mechanisms do not require continuity of neural tube tissue along the dorso-ventral axis and do not involve a sorting out of committed progenitors. Upon completion of theregeneration process, the pattern of neuron-specific transcription factor expression was essentially normal.Modest deficits in the numbers of transcription factor-defined neuron types were evident in the regeneratedtissue, increasing particularly in dorsal neuron types with later lesions. These results confirm the regulativepotential of the spinal neural tube and demonstrate a capacity for re-establishing appropriate cellular pat-terning despite a grossly abnormal morphogenetic situation.
Regulative regeneration is a well-known phenomenon in manyembryonic tissues including the developing nervous system. It in-volves a replacement of ablated embryonic cells through a compensa-tory proliferation and redistribution of other embryonic cells. It isoften referred to simply as regulation to distinguish it from wound
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healing and regeneration of adult tissues (Vaglia and Hall, 2000), al-though this definition does not require compensatory tissue growth.Early studies in amphibian and avian embryos using first classical em-bryological and histological techniques and later [3H]thymidinebirthdating and anterograde axonal tracing showed that most, if notall, regions of the neural tube have the capacity to repair restricted le-sions through compensatory proliferation in neighboring neural tis-sue (reviewed in Cowan and Finger, 1982). Although these earlystudies clearly demonstrated the regulative nature of the regenera-tion in terms of supplying an appropriate amount of tissue with nor-mal morphological and cytoarchitectonic appearance, there wasunderstandably little cellular and molecular characterization of theresultant regenerated tissue.
More recent studies, particularly focusing on the hindbrain neuraltube and using modern molecular anatomical techniques, haveshown that regulative regeneration generates tissue that is appropri-ately patterned (Diaz and Glover, 1996; Hunt et al., 1995; Jungbluth
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et al., 1997; Koster and Fraser, 2006; Saldivar et al., 1997; Scherson etal., 1993; Sechrist et al., 1995; Vaglia and Hall, 1999, 2000). Only oneof these studies (Diaz and Glover, 1996) investigated regulative re-generation of the entire dorsoventral extent of the neural tube, andshowed that selected neuron populations differentiated in their nor-mal locations. A comprehensive characterization of the patterning, in-cluding the dynamics of developmental patterning gene expression,was not obtained. Thus, how patterning is achieved in the face ofthe dramatic morphogenetic changes that take place during regula-tive regeneration has only been investigated superficially. Severalscenarios can be imagined. For example, extra committed progenitorsand/or neuronal precursors could be generated in the adjacent intacttissue and migrate to fill the lesion, sorting themselves out into theappropriate pattern while en route or after settling. Alternatively, un-committed progenitors or neuronal precursors could migrate to fillthe lesion, and become patterned thereafter by the same mechanismsextant in normal tissue.
A number of transcription factors have now been implicated in thepatterning of the spinal cord, including transcription factors that de-fine separate progenitor cell populations and transcription factorsthat are expressed specifically in the postmitotic neurons generatedby each progenitor cell population. Progenitor cell transcription factorexpression is triggered by gradients of signalling molecules emanat-ing from dorsal and ventral midline structures, and defines, throughcross-repressive interactions, sharply bounded dorsoventral domainswithin the proliferative zone (Briscoe and Novitch, 2008; Dessaud etal., 2008; Helms and Johnson, 2003). Each domain produces a cohortof neurons that expresses one or more specific transcription factors,giving rise to molecularly diverse neuron populations (Dasen, 2009;Goulding, 2009; Helms and Johnson, 2003). Increasing evidence indi-cates that the expression of postmitotic transcription factors is a cen-tral element in defining the identity of spinal neurons, including theirneurotransmitter profiles and axon trajectories (Goulding, 2009;Ladle et al., 2007). With this repertoire of relevant patterning genesthe process of regulative regeneration in the neural tube can nowbe characterized more comprehensively than was possible before.
Here, we provide new information about the capacity for regula-tive regeneration of that part of the neural tube that gives rise tothe lower thoracic and upper lumbar spinal cord in the chicken em-bryo. We characterize the patterning of the regenerated neural tissue
Fig. 1. Unilateral lesion of the spinal neural tube. A. Illustration of surgical procedure. Surfaceportion of which is removed unilaterally, sparing the floor plate. B, C. Transverse sections ofsal up. Arrow in C points to spared floor plate. D. Dorsal view of embryo immediately after
with respect to the expression of transcription factors that are in-volved in the differentiation of specific neuronal subtypes. We extendprevious work by showing that, like the hindbrain neural tube, thespinal neural tube has a pronounced capacity for regulative regener-ation. We show further that this regeneration is capable of recapitu-lating a normal pattern of gene expression and neuronaldifferentiation, although some deficits in neuronal numbers doarise, and that this patterning does not occur through a mixing andlater sorting out of progenitor subpopulations. Our results have rele-vance for attempts to use endogenous or exogenous stem and pro-genitor cells to regenerate adult neural tissue, where an importantgoal is to ensure proper patterns of neuronal generation anddifferentiation.
Materials and methods
In ovo surgery
White Leghorn chicken (Gallus gallus domesticus) eggs obtainedfrom local suppliers were incubated in a forced draft incubator at38–39 °C until stages HH14–17 (Hamburger and Hamilton, 1992).Surgical procedures (Fig. 1) were as described in Boulland et al.(2010). Portions of the neural tube tissue approximately 1–2 seg-ments long (one segment corresponding to the length of the adjacentsomite) were excised usually along with some of the overlying sur-face ectoderm, leaving the somites, floor plate and notochord intact.In sham controls, the procedure was carried out to the point of open-ing the vitelline membrane over the embryo, but no incisions weremade in embryonic tissue.
A total of 411+92 (for EdU labeling experiments)+53 (for multi-segment lesions)+30 (for DiI fate-mapping)=586 embryos wereused for histological and microscopic studies (an additional 161 em-bryos were used for Q-PCR experiments, see below). Four groups ofembryos received single segment neural tube lesions. In Group A(n=257+43 (for EdU labeling experiments)=300 embryos) lesionswere made at stage HH14–15 approximately at the level of somites24–25. In Group B (n=124 embryos) lesions were made at stagesHH16–17 approximately at the level of somites 24–25. In Group C(n=30 embryos) lesions were made at stages HH16–17 approxi-mately at the level of somites 16–17. In Group D (n=30 embryos),
ectoderm (blue) is cut away or deflected to reveal the underlying neural tube (green), athe neural tube prior to (B) and after (C) lesion, stained with haematoxylin–eosin. Dor-lesion. Scale bars: 50 μm (B, C), 200 μm (D).
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lesions were made at stages HH14–16 approximately at the level ofsomites 24–25, and remaining intact tissue either contralateral tothe lesion or immediately rostral to the lesion was immediately la-beled with the lipophilic tracer DiI to assess the origin of regeneratingcells (see below). The spinal neural tube at somites 24–25 and so-mites 16–17 become respectively the lower thoracic/upper lumbarand the upper thoracic segments of the spinal cord.
In some experiments (n=53 embryos) longer (up to 5-segmentstretches of the neural tube) or alternating (3 alternating single seg-ments) lesions were performed.
Embryos were allowed to develop for different durations. Embryosoperated at the lumbar level of the neural tube at earlier or later de-velopmental stages (Groups A and B) were allowed to develop post-operatively for 4, 8, 12, 24 or 36 h, whereas embryos operated atthe thoracic level at later stages (Group C) were allowed to developfor 36 h. Embryos in Group D were also allowed to develop for 36 h.At termination, the eggs were reopened and the embryos examinedin ovo with a stereomicroscope to assess the degree of regulative re-generation before being processed for histology, immunohistochem-istry or DiI-labeling.
Histology and immunohistochemistry
Embryos were removed from the eggs, fixed in 4% PFA for 30 minand processed for cryostat sectioning (12 μm thick transverse sec-tions) using conventional procedures, except for embryos in GroupD, which remained in fixative for examination as wholemounts (seebelow). Early (4–24 h postoperation) and late stage (36 h postopera-tion) neural tube sections were assessed for the expression of progen-itor and postmitotic neuronal transcription factors, respectively(Table 1). Visualization of the basement membrane surrounding theneural tube was with an antibody against laminin (Sigma). Nuclearstaining was with Hoechst 33342 (Invitrogen). Digital images werecaptured using a LSM5 scanning laser confocal microscope (Zeiss,Germany). Cell profiles labeled with antibodies to postmitotic neuro-nal transcription factors were counted using Image J software (NIH).
Table 1Antibodies used for immunohistochemistry and the cell population(s) they recognize.
Antigen Immunogen Source, catalogue or clone number
Pax7 Chicken PAX7 a.a. 352–523 DSHB — clone PAX7, provided by APax6 Chicken PAX6 a.a. 1–223 DSHB — clone PAX6, provided by ANkx2.2 Chicken Nkx2.2 DSHB— clone 74.5A5, provided byShh SHH-N DSHB — clone 5E1, provided by ThHB9 MNR2, C terminal — GST fusion protein
made in E. coliDSHB — clone 81.5 C10, provided bJessell
Islet1 C-terminal portion of rat Islet-1 DSHB— clone 39.4D5, provided by
Islet2 Islet 2, GST fusion from E. coli linkerregion of protein
DSHB— clone 51.4H9, provided by
Engrailed1 Chicken Engrail-1 DSHB — clone 4 G11, provided by Tand Susan Brenner-Morton
Lim1+2 Rat Lim2 DSHB — clone 4 F2, provided by Thand Susan Brenner-Morton
Brn3a Synthetic peptide conjugated to KLHderived from within residues 400 tothe C-terminus of mouse BRN3A
Abcam — ab23579
Pax2 Recombinant fragment, correspondingto mouse amino acids 188/385 of Pax2
Abcam — ab37129
Lim3 Lim 3, GST E. coli fusion protein DSHB — clone 67.4E12, provided bJessell
Lmx1b Chicken LMX, 6-his protein E. coliexpressed
DSHB— clone 50.5A5, provided by
Laminin Laminin isolated from the basementmembrane of Englebreth Holm–Swarm(EHS) mouse sarcoma
Sigma — L9393
GAM Cy3 Mouse IgG, Cy3-linked from goat GE Healthcare (Amersham) PA430GAR Cy3 Rabbit IgG, CY3-linked from goat GE Healthcare (Amersham) PA430
Assessment of cell origin using DiI-labeling
Immediately after lesion, a micropipette with a 10 μmdiameter tipopening was filled with a suspension of fine DiI particles (1,1′-diocta-decyl-3,3,3′,3′-tetramethylindocarbocyanine) made by diluting a sat-urated solution of DiI in 100% EtOH 1:10 in 20% w/v sucrose. The tip ofthe micropipette was placed either against the medial wall of the in-tact side opposite the lesion or against the caudal end of the intactrostral hemi-segment. Short pressure pulses were applied using aPV830 Pneumatic Picopump (WPI, USA) to repetitively depositsmall volumes of DiI particles until a substantial area of the exposedsurface of the intact tissue was noticeably pink. The eggs were thenresealed and returned to the incubator.
EdU labeling and assessment of the ratio of EdU-positive cells
Following a series of preliminary experiments using the thymidineanalog BrdU (5-bromo-2′-deoxyuridine), the thymidine analog EdU(5-ethynyl-2′-deoxyuridine, Invitrogen, 200 μl of 2.5 mg/mL) wasinjected adjacent to the neural tube lesion, between the chorion and vi-telline membrane, in Group A embryos (n=43) and stage-matchedsham operated embryos (n=49), 4 h prior to termination (immediate-ly after lesion for 4 h survival, 8 h postlesion for 12 h survival, and soon). Mortality is shown in Table 2. Embryos were removed, processedand sectioned as for immunohistochemistry. Sections were incubatedwith EdU-Click-iT reaction cocktail (modified from manufacturer's rec-ipe by further diluting Alexa 488 Fluor azide component 1:5) for 30 minat RT, washed briefly with 3% BSA in PBS, then with PBS (3×5 min),counterstained with Hoechst 33342 and mounted.
Imageswere taken at 488 nmexcitation for Alexa 488 (EdU-positivenuclei) and 343 nm excitation for Hoechst 33342 (all nuclei). Imageswere oversaturated to get a sharp border between labeled and non-labeled areas, pixels within cell nuclei profiles were counted indepen-dently in the Alexa 488 Fluor azide and Hoechst 33342 channels usingImage J software (NIH), and the ratio of pixel counts in the two channelswas calculated separately for the lesioned and intact sides of the neural
Host Dilution Cell population recognized
tsushi Kawakami Mouse 1:1 dP3–dP6 progenitor cellstsushi Kawakami Mouse 1:1 P0-PMN and dP5–dP6 progenitor cellsThomas M. Jessell Mouse 1:1 P3 progenitor cellsomas M. Jessell Mouse 1:1 Floor plate and notochordy Thomas M. Mouse 1:1 Somatic motoneurons
Thomas M. Jessell Mouse 1:1 dI3 interneurons, DRG neurons, somaticmotoneurons, sympathetic preganglionicneurons
Thomas M. Jessell Mouse 1:1 Somatic motoneurons
homas M. Jessell Mouse 1:1 v1 interneurons
omas M. Jessell Mouse 1:1 dI1, dI2, dI4 and dI6 interneurons
Rabbit 1:100 dI1–3 and dI5 interneurons
Rabbit 1:100 dI4, dI6, v0 and v1 interneurons
y Thomas M. Mouse 1:5(supernatant)
Motoneurons
Thomas M. Jessell Mouse 1:5(supernatant)
dI5 interneurons
Rabbit 1:100 Basement membrane/basal lamina
02 1:40004 1:400
Table 2Survival rates.
Single segment lesions (all groups) 406/441 92%Group A (lesion at HH14–15, somites 24–25) 242/257 94%Group B (lesion at HH16–17, somites 24–25) 112/124 90%Group C (lesion at HH16–17, somites 16–17) 24/30 80%Group D (lesion at HH14–16, somites 24–25) 28/30 93%Multiple segment lesions (all groups) 28/53 52%2 somites long 8/9 88%3 somites long 5/6 83%4 somites long 2/5 40%5 somites long 3/7 43%3×1 segment alternating 10/26 38%EdU-labeled embryos (with lesions) 38/43 88%Up to 24 h post-operative 35/35 100%36 h post-operative 3/8 37%EdU-labeled embryos (sham control) 44/49 89%Up to 24 h post-operative 41/41 100%36 h post-operative 3/8 37%
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tube. Ratios obtained by this method were the same as those calculatedfrom manual cell nuclei counts.
Real time quantitative PCR (Q-PCR)
All Q-PCR analyses were made on tissue from embryos lesioned(n=85) or sham-operated (n=76) at stages HH15–16. Embryosused for Q-PCR were in addition to the 507 embryos used for histolo-gy and microscopy described above. To increase the amount of regen-erating tissue available for Q-PCR analysis, multiple segment-longlesions were made in each lesioned embryo. Every other segmentwas lesioned unilaterally in the region encompassing somites 20 to24, leaving intact the segments adjacent to somites 19, 21, 23 and25, such that 3 ablated hemisegments were surrounded by 11 intacthemisegments. Small particles of charcoal were placed on the neuraltube adjacent to somite 18 to help target the tissue harvesting, whichwas done 12, 24 or 48 h post-lesion, after first categorizing the extentof regeneration (Table 3). Harvested tissue included all segments ad-jacent to somites 19 to 25, bilaterally. Tissue collection was doneunder RNAse-free conditions into RNAlater (Ambion, Applied Biosys-tems, USA). Two biological replicates were obtained for each timepoint, and tissue from sham and lesioned embryos was pooled sepa-rately (12 h lesioned: n=23 and 15; 12 h sham-operated: n=17and 24; 24 h lesioned: n=20 and 17; 24 h sham-operated: n=15and 12; 48 h lesioned: n=5 and 5; 48 h sham-operated: n=4 and 4).
Table 3Post-lesion mortality and degree and progress of regeneration.
Group Hours ofregeneration
Totalembryos
Died Noregeneration
Partrege
A 4 h 56 27 (48.2%) 29 (8 h 29 11 (37.9%) 18 (12 h 67 3 (4.5%) 12 (17.9%) 38 (24 h 42 1 (2.4%) 5 (11.9%) 24 (36 h 63 11 (17.5%) 5 (7.9%) 20 (
RNAwas purified using Total RNA Purification Kit (Norgen, Canada).DNAse (~0.35 Kunitz units/μl, RNase-Free DNase, Qiagen, Germany)was included to eliminate genomic DNA. RNA concentrations were de-termined usingNanoDrop (Thermo Scientific, USA) andRNAquality an-alyzed on a 2100 Bioanalyzer (Agilent Technologies, USA). Reactions of1 μg RNAwere set up for cDNA amplification using 2.5 μM random hex-ameres (Invitrogen, USA), 0.625 mM nucleotides (Promega, USA), 80 URNase OUT Recombinant Ribonuclease Inhibitor (Invitrogen, USA),5 mM DTT, 1× RT buffer and 400 U SuperscriptIII (Invitrogen, USA) ina total volume of 40 μl. RNA and random hexameres were incubatedat 65 °C for 5 min, then on ice for 1 min and at RT for 5 min. Mastermixof the other reagents was added and tubes were incubated at 50 °C for60 min and 70 °C for 15 min. The cDNA was then stored at −20 °Cprior to Q-PCR.
The Q-PCR reactions were run on calibrator (sham) and sample(lesion) cDNA from different times of regeneration (12, 24 and 48 hafter surgery) in 98-well Optical Reaction Plates and Optical AdhesiveCover (Applied Biosystems) using the ABI 7700 Sequence DetectionSystem (PE Applied Biosystems). Amplification reaction was 1× or2× SYBR Green PCR master mix (Applied Biosystems, USA), 1 μl ofcDNA and 400 nmol of forward and reverse primers in a total volumeof 20 μl. The PCR reaction had a standard amplification scheme: 1 cycleof 2 min (50 °C), 1 cycle of 10 min (95 °C), followed by 40 cycles of de-naturation for 15 s (95 °C) and annealing/extension for 1 min (60 °C).Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) and18S rRNAwere used as endogenous normalizers. All reactions were run in trip-licate. For the one biological replicate several plates were run to com-pare gene expression in sham and lesion cDNA, for the other genestested were located in a different pattern on the plates, providingsome randomization of the reactions to avoid systematic errors.
Sequences of genes suspected to be involved in regulative regen-eration were obtained from www.ensembl.org/Gallus_gallus andgene-specific primers were designed with Primer Express 3.0 soft-ware (Applied Biosystems, USA) using default parameters. Allprimers spanned exon–exon junctions, when this information wasavailable. All primers were validated by a template titration assayon a mix of cDNA from day (d) 2, d3, d4 and d5 chicken embryos. Di-lution series of cDNA reverse transcribed from RNA were set up (1 μg,0.2 μg, 0.1 μg, 0.01 μg and 0.001 μg). Controls were run in the absenceof template and in the absence of reverse transcriptase. Samples wererun with the amplification reaction and PCR scheme described aboveand for each gene a standard curve was plotted and the dissociationcurve analyzed to make sure it had a single peak at the amplification.Valid primers had a slope of about −3.3 and a correlation coefficient
122 G. Halasi et al. / Developmental Biology 365 (2012) 118–132
(R2-value) >0.95 for the curve (Applied Biosystems 2001, ABI Prism7900HT User Manual).
We used the comparative Ct method (or ΔCt) to find the relativechanges in mRNA levels between the calibrator (sham) and the sam-ple (lesion). Cycle times, rather than interpolated template quantities,were used for calculations. As required for the comparative Ct meth-od, the amplification efficiencies of compared genes were the same(Applied Biosystems 2001, ABI Prism 7900HT User Manual). Foreach of the 3 triplicates of a sample, the average cycle time (Ct), thestandard deviation (stdev), the coefficient of variation (Applied Bio-systems 2001, ABI Prism 7900HT User Manual), and the relative andfold changes in the sample (lesion) versus the calibrator (sham)were calculated (as described in Bookout and Mangelsdorf, 2003).
Statistics
Data were assessed using the Mann–Whitney U-test both manuallyand with GraphPad Prism software (GraphPad Software, Inc, La Jolla,CA, USA).
Fig. 2. Temporal progression of regulative regeneration. Transverse sections of the neural tuages (E–I, lesioned) at indicated hours post-lesion. Shown are examples of partial regeneraeration with a gap but with neural tube tissue present ventrally (12 h post-lesion, F, K, P), s(12 h post-lesion, G, L, Q, tissue bridge shown by arrows in G and L), complete regenerationR, granular appearance shown by arrows in H) and complete regeneration where the regenrowheads in E–I indicate anteroposterior levels of the sections in the indicated panels (J–R).and emphasize the gap between control and lesioned sides.
Results
General features of regulative regeneration in the spinal neural tube: sin-gle segment lesions
We grouped operated embryos according to the stage and the so-mite level at which the lesion was made. In Group A the lesion wasmade at HH14–15 and somite 24 or 25, in Group B the lesion wasmade at HH16–17 and somite 24 or 25 (2–10 h later than Group A),and in Group C the lesion was made at HH16–17 at somite 16 or 17(equivalent to 5–6 h later than Group B, given the rostrocaudal pro-gression of development in this region). As shown in Table 2, 92% ofthe embryos survived the surgery and developed to the chosen end-points (4–36 h post-operation). Development appeared to proceedat a normal rate after the operation. For example, somite number in-creased unabated, except for a moderate delay at 24 h (Supplementa-ry Fig. 1). General features of the regeneration were noted in 441embryos (257 in Group A, 124 in Group B, 30 in Group C, and 30 inGroup D).
be stained with haematoxylin–eosin (A–D, control; J–R, lesioned) and wholemount im-tion with a gap containing no neural tube tissue (8 h post-lesion, E, J, O), partial regen-lender rostral-to-caudal tissue bridges established along the lateral aspect of the lesionwhere the regenerated tissue has a granular appearance (“CRa”, 24 h post-lesion, H, M,erated tissue is indistinguishable from normal tissue (“CRb”, 36 h post-lesion, I, N). Ar-Scale bars: 50 μm (A–D, J-R); 200 μm (E–I). Figure H was retouched to darken the lumen
Fig. 3. Assessment of the origin of regenerated tissue. Wholemount images of 36 hregenerated neural tubes in which the opposite intact side (A, B) or adjacent rostral in-tact hemi-segment (C) was labeled with DiI immediately after lesion. Lesions weremade on the right sides of the neural tubes as seen in the images. Arrows indicateDiI-labeled tissue that has developed at the site of application. A) Example of a clearcontribution to the regenerated side from DiI-labeled tissue on the opposite intactside (although moderately deformed, this neural tube shows this contribution mostclearly of the examples we obtained). B) Example in which DiI-labeled tissue on theopposite intact side did not contribute to the regenerated side. An area lateral to theregenerated side has been masked to cover a distracting fluorescent artifact. C) Typicalexample of what was observed following DiI application to the adjacent rostral intacthemisegment. Little if any contribution to the regenerated tissue is seen (the faint fluo-rescence within the regenerating tissue is actually reflection of the strong fluorescenceof the application site). Scale bar 200 μm.
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Regeneration began rapidly. Minutes after the lesion, the dorsalaspect of the intact side reflected towards the adjacent somite, pre-sumably due to lateral tension exerted by the surface ectoderm. Theablated side contained at most a few scattered cells, and the ipsilater-al somite often bulged into the empty space (Figs. 2C and 3E and J). By4 h after lesion roughly 50–60% of the embryos (Groups A and B,Table 3) showed signs of partial regeneration, defined as a clear out-growth of neighboring intact tissue into the lesion (Figs. 2E, F, K, andP). This proportion increased to about 60–70% by 8 h; the remainderof embryos showed no signs of regeneration. Growth through thistimepoint had generated longitudinal extensions into the lesionfrom the adjacent intact ipsilateral hemisegments (Fig. 2F) and exten-sions primarily from the ventral aspect but in some cases also fromthe dorsal aspect into the lesion as seen in transverse sections(Figs. 2K and P). By 12 h after the lesion, growth had generated tissueextending well into the lesion from all 4 of these fronts and some em-bryos (about 20%) exhibited a slender bridge of tissue spanning thefull rostrocaudal and dorsoventral extent of the lesion (Table 3 andFigs. 2G, L, and Q). By 24 h in some embryos (about 10% in Group A)the lesion was completely filled with tissue that was continuouswith the adjacent neural tube, and we defined this as complete regen-eration. The appearance of this tissue varied, leading us to define twosubtypes of complete regeneration (CRa and CRb, Table 3). In CRa, thenew tissue had a granular appearance when obliquely illuminated, incontrast to the smooth appearance of the original neural tube tissue(Fig. 2H), as well as a different, yellower coloration (not shown, dueto difficulties capturing this on film). In CRb, the new tissue had thesame appearance as the original tissue (Fig. 2I). Since CRb firstappeared at 36 h, after CRa, we surmise that CRb represents a trans-formation from CRa through which the new tissue becomes more co-hesively organized. We observed no differences in the molecularcharacteristics of the regenerated tissue in CRa and CRb described inlater sections. The proportion of embryos exhibiting complete regen-eration (CRa or CRb) reached about 30% in Group A, 16% in Group B,13% in Group C and 20% in Group D by 36 h.
Origin of the regenerating tissue
Previously we used DiI labeling to show that the regenerating tis-sue that compensates for unilateral lesions of the hindbrain neuraltube originates both from the intact opposite side and from the
adjacent ipsilateral intact hemisegments (Diaz and Glover, 1996).Using the same approach here, we found that in the spinal neuraltube regenerating tissue clearly can derive from the opposite side,but we found no clear evidence of a contribution from the adjacentipsilateral hemisegments. In 30 embryos that were lesioned at aboutsomites 24–25 at stages HH14–16 (Group D), we applied DiI eitherto the opposite intact side (Group D1, n=16) or to the caudal endof the rostral adjacent intact hemisegment (Group D2, n=14). After36 h of regeneration, in Group D1 4 embryos exhibited complete, 6embryos nearly complete (tissue bridge), and 2 embryos partial re-generation, and in Group D2 2 embryos exhibited complete and 7 em-bryos partial regeneration (Table 3). Fig. 3 shows typical examples ofthe resultant DiI labeling. In half of the Group D1 embryos with com-plete regeneration, DiI-labeled patches of cells were clearly visiblewithin the regenerated tissue. In the other half, the regenerated tissuewas devoid of DiI-labeled cells. In these embryos, the regenerated tis-sue must have originated either from unlabeled portions of the intactopposite side or from the rostral and caudal adjacent intact hemiseg-ments. However, in Group D2 embryos the typical result was a re-stricted patch of DiI-labeled cells well behind the advancing tissuefront, corresponding roughly to the original DiI application site atthe edge of the lesion. In only one case did we see a few DiI-labeledcells within the regenerated tissue. From this we conclude that the in-tact opposite side definitely contributes to regeneration after unilat-eral lesions in the spinal neural tube, but that not all progenitors onthat side necessarily contribute. Furthermore, the obvious extensionof tissue from the rostral edge of the lesion is not accompanied by aclear contribution of cells from that location, but rather may repre-sent a gradual edge-to-midlesion filling in by cells from the oppositeside.
Regulative regeneration in the face of more extensive lesions
To further test the capacity of the spinal neural tube to regenerate,we made more extensive lesions of either 2–5 successive segments or3 alternating segments, in all cases unilaterally. Mortality over 24–26 h increased with amount of tissue ablated (Table 2), but in all sur-viving embryos regeneration occurred at all lesion sites. By 12 h afterlesion new tissue had appeared in all ablated segments in survivingembryos, and by 24 h the surviving embryos with 2-segment lesionsor 3 alternating-segment lesions exhibited either tissue bridges orCRa at all lesion sites (Fig. 4 and Table 4). For the larger lesions onlypartial regeneration was seen after 24 h (Table 4). Taking the ratioof remaining segments adjacent to the lesioned segments as a coarsemeasure of regenerative capacity, evidently once this ratio falls below3 the regenerative load exceeds capacity and the response is not ro-bust enough to regulate completely.
Regenerative capacity depends on developmental stage
To determine whether the neural tube at later stages can supportregulative regeneration, we first made lesions at somites 24–25 at thelater stage HH16–17 (Group B; neural tube development 2–10 h laterthan Group A). The frequency of complete regeneration (Table 3) by36 h postlesion in Group B was about 16% (8/49) compared to 30%(19/63) in Group A, with about 6% (3/49) of embryos exhibitingregenerated tissue indistinguishable from normal tissue comparedto about 12% (8/63) in Group A. Attempts at making lesions at so-mites 24–25 at even later stages became prohibitively difficult be-cause the neural tube became less accessible as the embryo turned.We therefore made lesions at somites 16–17 at HH16–17 (Group C),which based on the rostrocaudal gradient of development (approxi-mately 1.5 h per somite) would translate to about HH18–19 at so-mites 24–25 (7–16 h later in development compared to Group A).In this case the frequency of complete regeneration by 36 h postlesionwas only about 13% (4/30), with around 3% (1/30) of embryos
Fig. 4. Regeneration of multiple lesions. In ovo images of an embryo in which 3 one-hemisegment alternating lesions were made at stage HH14–15 adjacent to somites 21, 23 and 25(marked with arrowheads). Images were taken immediately (A), 12 h (B) and 24 h (C) post-lesion. Note that in this particular embryo chosen for sequential photography the al-ternating lesions were (atypically) shorter than one somite length. Scale bars: 200 μm.
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exhibiting regenerated tissue indistinguishable from normal tissue.These results indicate that regulative regeneration can occur at laterstages but becomes less robust as the neural tube matures.
Regeneration involves only modest increases in cell proliferation
To assess the degree of proliferation that accompanies regulativeregeneration, we used EdU labeling to obtain counts of dividingcells within different time epochs following the lesion (Figs. 5 and6). As a standard curve for proliferation, we labeled unoperated con-trol animals during the same time epochs. In preliminary experi-ments we used BrdU, which is visualized using antibodies, but wediscovered that EdU visualized with the much smaller molecular re-agents in the Click-iT reaction cocktail consistently gave on theorder of 50% more labeled nuclei than BrdU under identical condi-tions in the same stage embryos (not shown). Evidently this is simplya matter of superior access of reagents to the nucleus in the 12 μmthick sections we used.
As expected, the percentage of EdU-labeled cells during the last4 h of each time epoch gradually decreased with development, from60% at the 22–27 somite stage (4 h post-lesion, HH14–15) to 30% atthe 43–44 somite stage (36 h post-lesion, HH21–23) (Fig. 6). In le-sioned embryos, we found statistically significant increases in thepercentage of EdU-labeled cells at 4 h and 12 h after lesion (about10% and 5%, respectively) compared to controls, but no differencesthereafter (Fig. 6).
To corroborate this increase in proliferation, we performed Q-PCRfor a set of cell cycle-related genes on the tissue neighboring 3 alter-nating lesioned segments 12 h, 24 h and 48 h after lesion, and com-pared the mRNA levels to those in sham-operated controls.Upregulation of several cyclins and cyclin-dependent kinases (about20–50% relative to control), and downregulation of the cell cycleexit gene CEND1 (about 20% relative to control) was seen at 12 hpost-lesion (Fig. 7). By 24 h post-lesion, only one cyclin-dependentkinase (CDKN1) remained upregulated, and by 48 h cyclins and
Table 4Multiple lesion mortality and degree and progress of regeneration.
Lesion type Hours of regeneration Total embryos
2 continuous hemisegment 12 h 924 h 9
3 continuous hemisegment 12 h 624 h 6
4 continuous hemisegment 12 h 524 h 4
5 continuous hemisegment 12 h 724 h 7
3 alternating hemisegment 12 h 2624 h 17
cyclin-dependent kinase mRNA levels had largely normalized (onecyclin in fact showed about a 20% downregulation, Fig. 7). Cellcycle-related transcripts thus parallel the timecourse of regulative re-generation, but only show moderate changes.
To determine whether such modest increases in proliferationcould be sufficient to mount a compensatory proliferative response,we made a theoretical calculation based on the assumptions thatcell divisions during regulative regeneration are symmetrical andthat for increasing amounts of regenerated tissue required, cell pro-duction must be augmented by increasing the cell division rate(Appendix 1). Fig. 8 shows the increase in cell division rate requiredfor different numbers of intact hemisegments to compensate for onelesioned hemisegment, for any given number of cell cycles normallyoccurring during the time required for regulative regeneration toreach completion. Since the normal cell cycle time in the chicken em-bryo neural tube has been estimated to be 8 h (Langman et al., 1966),if we set 24 h as the time required for regulative regeneration andthus 3 as the number of cell cycles normally occurring during thattime window, we see that for the case of one lesioned hemisegmentsurrounded by 5 intact hemisegments, an increase of only 10% incell division rate is required for complete regeneration. This is verysimilar to the increase in proliferation seen using EdU labeling, andconceivably could be related to the observed moderate and transientincreases in cell cycle-related gene transcripts.
Progenitor transcription factor expression is recapitulated duringregeneration
During the initial stages of the regenerative response the base-ment membrane surrounding the neural tube was disrupted, some-times permitting surrounding mesenchymal cells to spread into thegap created by the lesion (Figs. 9A–C). Despite this, coherent tissuefronts advanced from the dorsal and ventral aspects of the unoper-ated side. An initial collapse of the dorsal and ventral aspects of the
Fig. 5. Detection of proliferating cells during regulative regeneration. EdU-labeling (green) of the nuclei of proliferating cells in regenerating (upper row) and intact (middle row)portions of the spinal neural tube of stage HH14–15 embryos with single-segment lesions adjacent to somite 24 or 25 and in stage-matched control embryos (lower row). EdU wasapplied 4 h prior to termination of the regeneration process at 4 h, 12 h, 24 h or 36 h postlesion. Lesioned side on right. Hoechst 33342 nuclear counterstaining in magenta. Nucleilabeled with both EdU and Hoechst 33342 are white. Scale bars: 50 μm.
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intact hemisegment towards the center of the lesion was sometimesobserved, but this never brought the two into apposition, and an ob-vious addition of tissue to each was typically observed by 12 h, oftenunequally such that one advancing front (most often the ventral) out-paced the other. By 12–24 h post-lesion the basement membrane wasfully re-established, and further regeneration occurred within it (Figs.9D–I). During this time, expression of the progenitor-specific tran-scription factors Pax7, Pax6 and Nkx2.2 and of the floor plate markerShh was evident within the advancing tissue fronts, and always main-tained the proper dorsoventral order even when regeneration waspartial and the regenerating tissue was discontinuous in the dorso-ventral axis (Figs. 9D, E, G, and H; Supplementary Fig. 2). TheNkx2.2 domain always extended immediately dorsal to the Shh do-main (sometimes overlapping it, see Figs. 9G and I), the Pax7 domain
Fig. 6. Quantitation of proliferation. Proliferation expressed as percentage of cells la-beled by EdU at different times post-lesion (4 h, 12 h, 24 h, and 36 h) in lesioned(open squares) and sham control (filled squares) neural tubes. EdU was applied 4 hprior to termination of regeneration at the indicated hours post-lesion. Differencestested statistically using the Mann–Whitney U-test (n=8 to 21, U=16.00 to 29.00,*p≤0.05).
was always the most dorsal, starting from the roof plate, and the Pax6domain always overlapped the ventral part of the Pax7 domain andstopped dorsal to the Nkx2.2 domain. Examination of partially regen-erated neural tubes showed that the overlaps and boundaries of theprogenitor domains maintained these relative positions regardlessof the actual transverse area of the tissue or the number of transcrip-tion factor-positive cells (Figs. 9D, E, G, and H). In some cases (Sup-plementary Figs. 2A and B) the three transcription factors wereexpressed within a continuous stretch of regenerating tissue on oneor the other side of the tissue gap, whereas in others (SupplementaryFigs. 2D and E) the tissue gap was located within one of the domainsor between two of them. There was never any sign of an initial abnor-mal mixing and later sorting out of cells expressing progenitor-specific transcription factors (the Pax6 and Pax7 domains normallyoverlap).
Normal progenitor and postmitotic transcription factor expression pat-terns are re-established in completely regenerated tissue
In embryos with complete or nearly complete regeneration theexpression patterns of the 3 progenitor-specific transcription factorsand of postmitotic transcription factors that define a broad set of dor-sal and ventral postmitotic neuron populations (Brn3a, Lim 1/2, Pax 2,Lmx1b, Engrailed 1, Islet 1/2, Lim 3 and HB9/MNR2) were completelyre-established and essentially indistinguishable from the normal pat-terns (represented by the control side, which did not differ fromsham-operated or unoperated embryos; Fig. 10). The numbers ofneurons in specific postmitotic neuron populations did not differ sig-nificantly on the regenerated and control sides of Group A embryos,although there was a slight trend towards lower numbers on theregenerated side (Fig. 11). In Group B and C embryos, on the otherhand, which received lesions at later developmental stages, the num-bers of neurons in certain dorsal postmitotic neuron populations (inGroup B), and in nearly all postmitotic neuron populations (inGroup C) were clearly lower on the regenerated side than on the con-trol side (Fig. 11). Thus, in parallel with the developmental decline ingeneral regenerative capacity described above, there is a
Fig. 7. Relative expression of indicated genes 12 h, 24 h and 48 h after lesion. Datapoints represent biological replicates, bars their means. Dashed lines indicate unity(no change in expression). Green points and bars represent cases where both replicatesexhibited consistent up- or down-regulation. Red points and bars represent cases withinconsistent replicates (only one replicate showed a change or the two replicatesshowed opposite changes). Black points and bars represent cases with no convincingchange in gene expression.
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developmental decline in the capacity to regulate the normal num-bers of different neuron types despite the ability to recapitulate nor-mal patterning, with dorsal neuron populations appearing to bemost sensitive.
Discussion
Regulative regeneration as a general phenomenon in the develop-ing central nervous system was described decades ago (Birge, 1959;Detwiler, 1944, 1946, 1947; Ferguson, 1957; Harrison, 1947;Holtzer, 1951; Kallen, 1955; Lehman and Youngs, 1952), but few at-tempts at molecular or cellular analysis have been made since(Buxton et al., 1997; Cowan and Finger, 1982; Diaz and Glover,1996; Hunt et al., 1995; Scherson et al., 1993; Sechrist et al., 1995).Our primary motivation here was to gain information about con-straints on the regenerative process and the degree to which theregenerated neural tissue becomes normally patterned and its con-stituent neurons differentiate. We have assessed molecular pattern-ing of the regenerating spinal neural tube by mapping theexpression of transcription factors that define different populationsof neuronal progenitors and postmitotic neurons. We have also ana-lyzed the capacity for regulative regeneration at different stages andwith varying types and sizes of lesions, and evaluated the extent towhich cell proliferation increases during regeneration. Our principalfindings are as follows: 1) the lower thoracic/upper lumbar spinalneural tube has the capacity to regenerate regulatively within limitsimposed by the regenerative load (ratio of ablated/remaining tissue)and the developmental stage of the embryo, 2) the regenerated tissuederives primarily from the opposite intact side, 3) successful regener-ation requires only a modest increase in cell proliferation rate, 4) pro-genitor patterning is recapitulated during regeneration without signsof mixing and later sorting out of progenitor subpopulations, and 5) anormal pattern of postmitotic neurons is re-established in the regen-erated tissue, but with numerical deficits following regeneration atlater developmental stages. Our analysis provides a better foundationfor investigating molecular mechanisms that underlie regulative re-generation in the neural tube.
Capacity for and limits to regeneration
Early studies reported a high capacity for regulative regenerationof the neural tube in amphibians, but variable results in the chickenembryo. We find that complete regulative regeneration of the spinalneural tube in chicken embryos is obtained in 15–30% of embryoswithin 36 h (depending on lesion size), similar to what has beenseen in the hindbrain neural tube (Buxton et al., 1997; Diaz andGlover, 1996; Scherson et al., 1993). The incidence of complete regen-eration declines as lesions are made at later developmental stages orencompass larger fields of tissue. Several factors probably contributeto limiting the success rate, including changes in the mechanical situ-ation and a progressive decline in proliferative capacity as progeni-tors are depleted.
The mechanical situation is probably very important but hard tocontrol experimentally. Clark and Scothorne (1990) showed thatwhen performed at early enough stages incision of the roof platealone does not result in splaying of the dorsal margins of the neuraltube, thereby permitting rapid healing. This may reflect the normalocclusion of the neural tube lumen, which in chick embryos starts atstage 10/11 and ends at stage 14/15 (Desmond and Schoenwolf,1985, 1986). Occlusion appears to be mediated by adhesive forces be-tween the two lateral lumen walls, and would tend to minimizesplaying. In embryos older than stage 14/15 the adhesive forces evi-dently decline rapidly, and splaying occurs more readily. This wasclearly the case in our experiments.
To assess the temporal dependence of regulative regeneration, wemade lesions at the same segmental level but at later stages (corre-sponding to 2–10 h further development) or at a more rostral, develop-mentally advanced segmental level (representing an additional 5–6 h offurther development). Because the rate of complete regeneration de-clinedmarkedlywith bothways of obtaining later lesions, it seems likelythat developmental stage is an important factor. What changes over
Fig. 8. Mathematical calculation of dependence of proliferative compensation on regenerative load. Each curve indicates how the augmentation of a given initial cell division ratevaries with the factor Z, which is related to the ratio of spared hemisegments (S) adjacent to lesioned hemisegments (m). See Appendix A for the mathematical formula used. Thelower the initial cell division rate, the greater the augmentation required to compensate for tissue loss.
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time might be responsible for limiting the regenerative response? Inprinciple this must be due either to a decrease in proliferative capacityor in the capacity to translocate from the intact neural tube or both.That proliferation in the spinal neural tube diminishes over time is wellknown (this can be seen in Fig. 6 in the unoperated control embryos),so a decrease in proliferative capacity seems likely to be a primary factor.Whether this is intrinsic or imposed by extrinsic factors needs to betested.
Origin of the regenerated tissue
The appearance of the regenerating tissue, which advances fromrostral, caudal, dorsal and ventral fronts, suggests that progenitorson all sides contribute to filling in the lesion. However, this could bean illusion, since rostral and caudal filling could in principal derivesolely from the opposite side (if cells first advanced from there intothe lesion along its rostral and caudal margins), and dorsal and ven-tral filling could in principal derive solely from the rostral and caudalintact hemisegments (if cells first advanced from these into the lesionalong its dorsal and ventral margins). Only fate-mapping or dynamicimaging can provide definitive evidence for tissue origins. In an earli-er study of regulative regeneration in the hindbrain neural tube, weused fate-mapping with DiI to show that both the intact oppositeside and the adjacent rostral and caudal ipsilateral hemirhombo-meres contribute to regeneration of a lesioned hemi-rhombomere(Diaz and Glover, 1996). Here, we find using the same approachthat the intact opposite side definitely contributes to regenerationof a lesioned hemisegment of the spinal neural tube. However, be-cause we only labeled a portion of the opposite side with DiI in eachneural tube, and only about half of regenerated hemisegments con-tained DiI-labeled cells, we must conclude that not all progenitorcells on the intact opposite side necessarily contribute to the regener-ation. Moreover, although we found no contribution from DiI-labeledintact rostral hemisegments here, we cannot rule out that such con-tribution never occurs, nor can we rule out a contribution from the in-tact caudal hemisegment, which we did not label. It seems likely that,as in the hindbrain, regenerating tissue can derive from all possibleadjacent sources of progenitor cells, but that the relative contribu-tions may vary greatly from case to case. The most effective way tocharacterize this potentially highly variable dynamic process wouldbe by sequential confocal live imaging after selectively labeling the
three sources with different fluorescent markers, such as geneticallyencoded fluorescent proteins. This is technically possible in the chick-en embryo using in ovo electroporation, and we suggest this as thelogical next step in pursuing this question.
Cell proliferation associated with regulative regeneration
We observed about a 10% increase in the cell division rate by 4 hpost-lesion, which fell to 5% by 12 h and was no longer evident by24 h. As corroboration, we found an upregulation of the transcriptionof several proliferation-related genes through 12 and 24 h post-lesion, and a down-regulation of a cell cycle exit protein (Fig. 7).Cowan and Finger (1982) also observed an increase in cell prolifera-tion (incidence of mitotic figures) in the regenerating mesencephalictectum of the chicken embryo, and Diaz and Glover (1996) observedan increase in cell number within the regenerating tissue in the hind-brain of the chicken embryo. Since the normal cell cycle time in thechicken embryo spinal neural tube has been estimated to be 8 h(Langman et al., 1966), we can estimate how much a 10% increasein proliferation would contribute to the regeneration of a single neu-ral tube hemisegment (Fig. 8). If we set 24 h and thus 3 cell cycles asthe time encompassed by regulative regeneration, we see that for thecase of one lesioned hemisegments surrounded by 5 intact hemiseg-ments, an increase of only 10% in cell division rate is required to attaincomplete regeneration. Conversely, if the cell cycle time is 12 h or theregenerative load becomes too great (Z>1.3), then a 10% increase inproliferation rate cannot compensate for the loss. Assuming cycletime is not increasing during the time window of the regenerative re-sponse, then the observed inability to compensate for the larger mul-tiple segment lesions is likely the result of exceeding the regenerativeload that the neural tube can tolerate.
In contrast to our findings, Buxton et al. (1997) concluded thatproliferation was not necessary for regeneration, since they observedregulation with a reduced neural tube diameter, and Scherson et al.(1993) reported that they saw no dramatic increase in cell divisionrate using BrdU incorporation. However, since we found in extensivepreliminary experiments that detection of newly mitotically activecell nuclei with immunohistochemistry for BrdU is markedly inferiorto EdU labeling with Click-iT technology, it is possible that the modestincrease we observed may have escaped the attention of Scherson etal. (1993) for this reason.
Fig. 9. Basal membrane regeneration and recapitulation of progenitor transcription factor expression patterns during regulative regeneration. A) Loss of basal membrane integrityrevealed by immunohistochemistry for laminin (red); nuclear counterstaining with Hoechst 33342 (blue). B,C) Regenerating neural tubes 12 h and 24 h post-lesion labeled withEdU (green) and Hoechst (magenta). Nuclei labeled with both are white. In A–C surrounding mesenchyme has entered the lumen (arrows) through the interrupted basal mem-brane. D–I) Basal membrane and transcription factor expression revealed by immunohistochemistry for laminin (red), Pax7 (magenta, dorsal domain), Pax6 (green), Nkx2.2 (ma-genta, ventral domain), and Shh (yellow). Overlapping colors in the images yield white, but do not necessarily represent double-labeled nuclei. Transverse sections, lesioned side onright, dorsal up. F and I show stage-matched sham controls. Continuity of the basal membrane is re-established at about 12 h post-lesion. The progenitor domains are compressedwithin the regenerating tissue while maintaining proper spatial relationships, and approach symmetry with the unoperated side as regeneration nears completion. Panels A and Dare of the same section. Scale bars: 50 μm (same scale bar for A,B, same scale bar for D–I).
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One of the most interesting questions regarding regulative regen-eration is how the regenerative response is terminated, and relatedly,why it fails in a large proportion of cases. We have previously sug-gested that one reason for premature termination might be the pres-ence of proliferating lateral mesenchyme that may enter the lesionand compete for space as the neural tube attempts to regulate (Diazand Glover, 1996; see Fig. 9). We have not made a systematic assess-ment of the presence of ectopic lateral mesenchyme in cases of partialor failed regeneration, but we note that there is typically no overt signof foreign tissue within the neural tube when this occurs. Of course,ectopic mesenchyme could create a temporary impingement that issuccessfully removed but sufficiently disruptive to prevent regenera-tion from proceeding to completion.
A general difficulty in addressing the question of premature termi-nation is that one cannot predict which neural tubes will fail to regen-erate, since failed regenerators look like successful regenerators thataren't yet finished. One must wait until successful regeneration iscompleted before assessing which neural tubes have failed. This prob-lem could be addressed by sequential live imaging as noted above.Nevertheless, our results already bear to some extent on the question,
in that we have made observations of proliferation and patterning atvarious time points prior to complete regeneration. Since the majorityof neural tubes do not regenerate completely, most of our observa-tions must stem from neural tubes that eventually fail. It is thereforelikely that in cases of failed regeneration, as in successful regenera-tion, there is a modest proliferative response and a recapitulation ofpatterning, up to the point of failure.
Progenitor patterning
The expression patterns of progenitor-specific transcription fac-tors during regeneration are particularly interesting and have beenperceived in previous studies as a major unanswered question re-garding the issue of whether regulative regeneration recapitulatesoriginal developmental patterns (Vaglia and Hall, 1999). However, itis also difficult to analyze quantitatively due to the dynamic and var-iable morphology of the regenerating tissue. As a consequence, ouranalysis of progenitor patterning is only qualitative. We note that atall stages of the regenerative response the three progenitor-specifictranscription factors Nkx2.2, Pax6 and Pax7 occupy domains in the
Fig. 10. Re-establishment of the expression patterns of selected postmitotic transcrip-tion factors. Transcription factors color-coded: Brn3a (green), Pax2 (magenta) and HB9(yellow) in A,C,E; Lim1/2 (red) and Islet1 (cyan) in B,D,F. Lmx1b, Engrailed 1 and Lim 3were also assessed qualitatively (quantitatively in Fig. 11) but not shown here. Stage-matched controls (A,B), complete regeneration 36 h post-lesion (C,D) and partial re-generation 36 h post-lesion (E,F). Transverse sections, lesioned side on right, dorsalup. Note the lower number of dorsal neurons (Brn3a, Pax2, Lim 1/2) on the regener-ated side (right) in incompletely regenerated neural tubes (E,F). Scale bars: 50 μm(same scale bar for A–D, same scale bar for E,F).
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proper ventral to dorsal order and of appropriately relative sizesroughly scaled to the amount of tissue present. This suggests the ac-tivity of the same signalling gradients and cross-repressive interac-tions that act on intact tissue (Briscoe and Ericson, 2001; Briscoeand Novitch, 2008), as well as a compression or size regulation ofthe patterning that has also been reported in the chicken embryohindbrain (Buxton et al., 1997). Most importantly, we have neverseen abnormal intermingling of cells expressing these transcriptionfactors. Even when little regenerated tissue is present, expression isin systematic dorsoventral domains, indicating that immigrating pro-genitors are not bringing their previously specified identities withthem from their origins in the adjacent intact tissue, but rather arebeing patterned by the normal signalling gradients on the regenerat-ing side. Otherwise we would have to postulate a highly regulatedmigration wherein the different progenitor subpopulations enterthe regenerating fronts in waves according to their ultimate dorso-ventral positions. Since we see proper patterning in cases where allthree progenitor subpopulations are present within either a singledorsal front or a single ventral front (Supplementary Fig. 2), such co-ordinated migration in successive waves would involve opposite se-quences for the two cases (Nkx2.2 followed by Pax6 followed by
Pax7 for the dorsal front, and the reverse for the ventral front). A mi-gratory choreography of this precision, though formally possible, be-gins to stretch the imagination. Our preferred interpretation, thatpatterning is imposed anew as the progenitors enter the regeneratingtissue, is reminiscent of the corrective respecification of Hox gene ex-pression in hindbrain progenitor cells that become mispositionedalong the anteroposterior axis (Trainor and Krumlauf, 2000).
We also note that the systematic deployment of the three tran-scription factor domains within separated dorsal and ventral frontsindicates that patterning can occur despite a tissue discontinuity. Al-though the dorsal and ventral signalling gradients are known to mu-tually antagonize each other (Dessaud et al., 2008; Ulloa and Briscoe,2007), and the influence of the dorsal gradient reaches all the way tothe ventral part of the neural tube (Ulloa and Marti, 2010; Yu et al.,2008), it is not clear to what extent the two gradients must interactto create a proper pattern. If an interaction is essential, then duringregulative regeneration the gradients would somehow have to beconducted across the gap. Diffusion of high molecular weight Shhcomplexes evidently occurs in part along the luminal surface(Chamberlain et al., 2008). It is unclear what route would be takenduring regeneration, when much of this surface has been disrupted.
We have not here examined the expression of transcription factorssuch as the HoxC proteins that demarcate anteroposterior domainsalong the thoracolumbar spinal cord. This may also be relevant giventhat we have not ruled out contributions to the regenerating tissuefrom the rostral and caudal intact hemisegments and thus longitudi-nal movement. We intend to address this issue in future studies.
Patterning of postmitotic neurons
Only a few studies have assessed neuronal differentiation withinthe regenerated neural tube, with focus on laminar organization ofneurons and inputs in the mesencephalic tectum (Cowan andFinger, 1982), the location of cranial nerve exit points (Buxton et al.,1997) and the location and numbers of specific hindbrain neurontypes, for which a numerical deficit was seen in the one dorsal neurontype assessed (Diaz and Glover, 1996). Here we obtain a more com-prehensive picture, using a large panel of transcription factors thatdefine molecularly distinct neuron subclasses. The neuron subclassesare located in the appropriate locations in the mantle zone followingregeneration, indicating that their parent progenitor populationshave been patterned sufficiently well to produce an essentially nor-mal distribution of neurons. When regeneration is forced to occur atlater developmental stages however, numerical deficits in all neuronpopulations start to appear, most pronounced in the most dorsal neu-ron types.
The bias in numerical deficit might be caused by a specific decre-ment in either dorsal proliferation or dorsal patterning or both. A dec-rement in proliferation could occur due to a loss of dorsally-derivedBMPs and Wnts, which are both mitogenic (Ulloa and Briscoe,2007). A decrement in patterning seems likely to be due to a deficientdorsal patterning per se as opposed to an encroachment of ventralpatterning, since we found no proportional increase in ventral neurontypes. Scherson et al. (1993) observed generally normal developmentexcept for deficiencies in dorsal structures following unilateral abla-tion of the mesencephalic and metencephalic neural tube, and hy-pothesized that this might be caused by ineffective signallinginteractions between the neural tube and the overlying surface ecto-derm. The expression of slug and Pax3 (and presumably Pax7) in thedorsal neural tube require this interaction, for example (Buxton et al.,1997). We did not specifically assess the spatial relationship betweenneural tube and surface ectoderm, but since we typically saw expres-sion of Pax7 dorsally during the regenerative process, there is no ob-vious indication from our experiments of deficient signalling betweenthem. What causes the more pronounced numerical deficits in dorsalneuron populations therefore remains an intriguing question.
Fig. 11. Counts of neurons expressing postmitotic transcription factors 36 h post-lesion. Average number of neurons in the regenerated (white columns) and ipsilateral adjacentrostral and caudal hemisegments (grey columns), each normalized to the number in the contralateral control (intact) hemisegment. Grand averages and s.d. across animals(n=3 to n=6) of average counts per section from at least 15 sections per animal. Differences were tested statistically using the Mann–Whitney U-test (n=4 to 6, U=0.00 to4.00, *p≤0.05, **≤0.01, ***≤0.001). Differences for Group B could not be tested because nb4.
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General conclusion
We conclude that regulative regeneration at early stages of neuraltube development in the chicken embryo involves a swift increase inproliferation that, although modest, is sufficient to compensate forthe loss of tissue. Neuronal patterning is robust despite an abnormalmorphogenetic context. Numerical deficits in neuron populationsthat occur with regeneration at later stages are probably linked to ageneral decline in regenerative potential. However, because they aremore pronounced dorsally, there may be a particular sensitivity of
dorsal neural tube to the regenerative situation, manifested eitheras deficient proliferation or patterning or both.
We feel that this information has a significance beyond the prov-ince of developmental mechanisms. With the use of fetal neural pro-genitors and embryonic stem cells being considered as possibletherapies for brain and spinal cord injury, a more comprehensiveknowledge of the potential (and limits to potential) of specific re-gions of the neural tube to proliferate and to generate appropriateneuronal populations will benefit the design of stem cell-based re-placement strategies.
131G. Halasi et al. / Developmental Biology 365 (2012) 118–132
Supplementary materials related to this article can be found on-line at doi:10.1016/j.ydbio.2012.02.012.
Acknowledgments
We gratefully acknowledge the technical assistance of Melinda Lil-lesand, Åse-Marit Kristiansen and Kobra Sultani. This study is sup-ported by the Norwegian Research Council Stem Cell Program, theNorwegian Center for Stem Cell Research, and postdoctoral fellow-ships to GH from the Norway-Hungary Cultural Exchange Programand AMS from the Norwegian Research Council. We thank James Bris-coe for critically reviewing the manuscript.
Appendix 1. Theoretical calculation relating compensatory in-crease in cell division rate to regenerative load.
We assume a symmetrical mode of cell division, such thatNC=N0•2C.
where
C=number of cell cyclesN0=original number of cellsNC=number of cells after a given number of cell cycles C
We assume further that to compensate for a loss of tissue, thenumber of cell cycles must increase within the same time windowby some factor X, such that
NCX ¼ N0•2CX
Where
NCX=number of cells after the augmented number of cell cyclesCX.
If we assume a single hemisegment ablation, and we allow thenumber of intact hemisegments S that contributes to the regenera-tion of the ablated hemisegment to vary, we can factor in S thus:
Sþ 1ð Þ•NC ¼ S•NCX
or
Sþ 1ð Þ•NC ¼ S•N0•2CX
such that following an ablation, S hemisegments generating tissue atthe augmented cell division rate produce the same amount of tissuethat S+1 hemisegments would generate at the normal division rate.
Similarly, if we assume multiple hemisegment ablations wherethe number of ablated hemisegments=m, and we allow the numberof intact hemisegments S that contributes to the regeneration of theablated hemisegment to vary, we can factor in S thus:
Sþmð Þ•NC ¼ S•N0•2CX
But NC ¼ N0•2C ¼ N0•Y
So Sþmð Þ•N0•2C ¼ S•N0•2
CX
Factoring out N0 we get
Sþmð Þ•2C ¼ S•2CX
Sþmð Þ=Sð Þ•2C ¼ 2CX
We introduce a new variable Z=((S+m)/S), so that
Z•2C ¼ 2CX
We can calculate X thus:
log10 Z•2C� �
¼ X•log10 2C� �
X ¼ log10 Z•2C� �
=log10 2C� �
Z is related to the ratio of intact (S) to ablated (m) hemisegments (in-deed, S/m=(S+m)/mZ), so plotting X versus Z as a function of C al-lows us to see how the increase in cell division relates to theregenerative load S/m, given different values of C (cell cycles normal-ly occurring during the time window of regulative regeneration) as aninitial condition (see Fig. 6).
References
Birge, W.J., 1959. An analysis of differentiation and regulation in the mesencephalon ofthe chick embryo. Am. J. Anat. 104, 431–463.
Boulland, J.L., Halasi, G., Kasumacic, N., Glover, J.C., 2010. Xenotransplantation ofhuman stem cells into the chicken embryo. J. Vis. Exp.
Briscoe, J., Ericson, J., 2001. Specification of neuronal fates in the ventral neural tube.Curr. Opin. Neurobiol. 11, 43–49.
Briscoe, J., Novitch, B.G., 2008. Regulatory pathways linking progenitor patterning, cellfates and neurogenesis in the ventral neural tube. Philos. Trans. R. Soc. Lond. B Biol.Sci. 363, 57–70.
Buxton, P., Hunt, P., Ferretti, P., Thorogood, P., 1997. A role for midline closure in thereestablishment of dorsoventral pattern following dorsal hindbrain ablation. Dev.Biol. 183, 150–165.
Chamberlain, C.E., Jeong, J., Guo, C., Allen, B.L., McMahon, A.P., 2008. Notochord-derivedShh concentrates in close association with the apically positioned basal body inneural target cells and forms a dynamic gradient during neural patterning. Devel-opment 135, 1097–1106.
Clark, B.J., Scothorne, R.J., 1990. Variation in the response of chick embryos to incisionof the roof plate of the neural tube at different developmental stages. J. Anat. 168,167–184.
Cowan, W.M., Finger, T.E., 1982. Regeneration and regulation in the developing centralnervous system. In: Spitzer, N.C. (Ed.), Neuronal Development. Plenum Press, NewYork, pp. 277–415.
Dasen, J.S., 2009. Transcriptional networks in the early development of sensory-motorcircuits. Curr. Top. Dev. Biol. 87, 119–148.
Desmond, M.E., Schoenwolf, G.C., 1985. Timing and positioning of occlusion of the spi-nal neurocele in the chick embryo. J. Comp. Neurol. 235, 479–487.
Desmond, M.E., Schoenwolf, G.C., 1986. Evaluation of the roles of intrinsic and extrinsicfactors in occlusion of the spinal neurocoel during rapid brain enlargement in thechick embryo. J. Embryol. Exp. Morphol. 97, 25–46.
Dessaud, E., McMahon, A.P., Briscoe, J., 2008. Pattern formation in the vertebrate neuraltube: a sonic hedgehog morphogen-regulated transcriptional network. Develop-ment 135, 2489–2503.
Detwiler, S.R., 1944. Restitution of the medulla following unilateral excision in the em-bryo. J. Exp. Zool. 96, 129–142.
Detwiler, S.R., 1946. A quantitative study of locomotion in larval Amblystoma followingeither midbrain or forebrain excision. J. Exp. Zool. 102, 321–332.
Detwiler, S.R., 1947. Restitution of the brachial region of the cord following unilateralexcision in the embryo. J. Exp. Zool. 104, 53–68.
Diaz, C., Glover, J.C., 1996. Appropriate pattern formation following regulative regener-ation in the hindbrain neural tube. Development 122, 3095–3105.
Ferguson, T., 1957. The restitution of the medulla oblongata following unilateral exci-sion in embryos of the frog. J. Exp. Zool. 135, 403–423.
Goulding, M., 2009. Circuits controlling vertebrate locomotion: moving in a new direc-tion. Nat. Rev. Neurosci. 10, 507–518.
Hamburger, V., Hamilton, H.L., 1992. A series of normal stages in the development ofthe chick embryo. 1951. Dev. Dyn. 195, 231–272.
Harrison, R.G., 1947. Wound healing and reconstitution of the central nervous systemof the amphibian embryo after removal of parts of the neural plate. J. Exp. Zool.106, 27–83.
132 G. Halasi et al. / Developmental Biology 365 (2012) 118–132
Kallen, B., 1955. Regeneration in the hindbrain of neural tube stages of chick embryos.Anat. Rec. 123, 169–181.
Koster, R.W., Fraser, S.E., 2006. FGF signaling mediates regeneration of the differentiat-ing cerebellum through repatterning of the anterior hindbrain and reinitiation ofneuronal migration. J. Neurosci. 26, 7293–7304.
Ladle, D.R., Pecho-Vrieseling, E., Arber, S., 2007. Assembly of motor circuits in the spinalcord: driven to function by genetic and experience-dependent mechanisms. Neu-ron 56, 270–283.
Langman, J., Guerrant, R.L., Freeman, B.G., 1966. Behavior of neuro-epithelial cells dur-ing closure of the neural tube. J. Comp. Neurol. 127, 399–411.
Lehman, H.E., Youngs, L.M., 1952. An analysis of regulation in the amphibian neuralcrest. J. Exp. Zool. 121, 419–447.
Saldivar, J.R., Sechrist, J.W., Krull, C.E., Ruffins, S., Bronner-Fraser, M., 1997. Dorsal hind-brain ablation results in rerouting of neural crest migration and changes in geneexpression, but normal hyoid development. Development 124, 2729–2739.
Scherson, T., Serbedzija, G., Fraser, S., Bronner-Fraser, M., 1993. Regulative capacity ofthe cranial neural tube to form neural crest. Development 118, 1049–1062.
Sechrist, J., Nieto, M.A., Zamanian, R.T., Bronner-Fraser, M., 1995. Regulative responseof the cranial neural tube after neural fold ablation: spatiotemporal nature ofneural crest regeneration and up-regulation of Slug. Development 121,4103–4115.
Trainor, P.A., Krumlauf, R., 2000. Patterning the cranial neural crest: hindbrain segmen-tation and Hox gene plasticity. Nat. Rev. Neurosci. 1, 116–124.
Ulloa, F., Briscoe, J., 2007. Morphogens and the control of cell proliferation and pattern-ing in the spinal cord. Cell Cycle 6, 2640–2649.
Ulloa, F., Marti, E., 2010. Wnt won the war: antagonistic role of Wnt over Shh controlsdorso-ventral patterning of the vertebrate neural tube. Dev. Dyn. 239, 69–76.
Vaglia, J.L., Hall, B.K., 1999. Regulation of neural crest cell populations: occurrence, dis-tribution and underlying mechanisms. Int. J. Dev. Biol. 43, 95–110.
Vaglia, J.L., Hall, B.K., 2000. Patterns of migration and regulation of trunk neural crestcells in zebrafish (Danio rerio). Int. J. Dev. Biol. 44, 867–881.
Yu, W., McDonnell, K., Taketo, M.M., Bai, C.B., 2008. Wnt signaling determines ventralspinal cord cell fates in a time-dependent manner. Development 135,3687–3696.
