Regeneration of Respiratory Pathways within Spinal Peripheral Nerve Grafts

Regeneration of Respiratory Pathways within SpinalPeripheral Nerve Grafts

PATRICK DECHERCHI, NATHALIE LAMMARI-BARREAULT, AND PATRICK GAUTHIER

Departement de Physiologie et de Neurophysiologie, Laboratoire de Biologie des Rythmes et du Developpement, URA CNRS 1832, GroupeNeurobiologie fonctionnelle des greffes nerveuses, Faculte des Sciences et des Techniques de Saint-Jerome (Aix-Marseille III),

Case Courrier 332-351-352, Avenue Escadrille Normandie Niemen, 13397 Marseille Cedex 20, France

Central respiratory neurons exhibit normal activityafter axonal regeneration within blind-ended periph-eral nerve grafts (PNGs) inserted near the correspond-ing cell bodies in the medullary respiratory centers.Part of these medullary respiratory neurons projecttoward the spinal cord and contribute to descendingrespiratory pathways that control respiratory moto-neurons. The present work investigates to what extentcervical respiratory pathways could be directed out ofthe central nervous system within PNGs inserted dis-tant to the medullary respiratory nuclei. In adult rats(n 5 13), autologous segments of the peroneal nervewere implanted into the ventrolateral part of the C2spinal cord at the level of the descending respiratorypathways. Two to four months after grafting, electro-physiological recording of teased graft filaments(n 5 562) revealed the presence of regenerated nervefibers with unitary impulse traffic (n 5 164) in alltested PNGs (n 5 6). Respiratory discharges (n 5 52)corresponded to efferent and afferent activity. Effer-ent respiratory discharges (n 5 32) originated fromcentral respiratory neurons which remained func-tional and preserved afferent connections. Retrogradehorseradish peroxidase labeling applied to the distalcut end of PNGs (n 5 7) revealed stained (42/1997)neurons in areas where respiratory cells have beendescribed. Afferent respiratory discharges (n 5 20)were synchronized with lung inflation but their origin(stretch pulmonary receptors and/or respiratorymuscle receptors) was not determined. On the basis ofadditional data from light and electron microscopy ofPNGs, comparison was made between anatomical, ret-rograde labeling, and electrophysiological data. Themain conclusion is that spinal PNGs appear to be ableto promote axonal regeneration of functional respira-tory efferent and afferent pathways. r 1996 Academic Press,

Inc.

INTRODUCTION

In contrast to the abortive regrowth that occurs whenaxons are interrupted in the adult mammalian central

nervous system (CNS) (6, 48), exposure of injured CNSaxons to the nonneuronal milieu of a peripheral nervecan lead to extensive axonal elongation (1, 5, 29, 48).The ability of different types of mature CNS neurons toregenerate their axons within peripheral nerve grafts(PNGs) is now clearly established (1, 2, 4, 5). Anadditional example of the remarkable plasticity of theadult mammalian CNS neurons is to be found in themodel of axonal regeneration of respiratory cells. Witha view to assessing the extent to which the functionalcapacities of CNS neurons can be recovered afteraxonal regeneration, we have examined the electro-physiological activity of respiratory neurons followingregeneration of their axons into blind-ended graftsimplanted within the medulla oblongata of the rat (23,24, 33, 34). Respiratory neurons regenerated theiraxons for distances of several centimeters into medul-lary PNGs and their functional state appeared similarto that of intact cells. Recordings of unitary activitiesfrom small filaments teased from the graft 2 to 5months after grafting showed that they retained nor-mal patterns of spontaneous respiratory activity andnormal responses to their usual inputs.These functional properties were established exclu-

sively using grafts inserted near the cell bodies, i.e., inthe respiratory centers of the medulla oblongata. How-ever, one major question that arises concerns theability of respiratory neurons to regenerate axonswithin PNGs inserted at some distance from the cellbodies. This is important because the distance from cellbody to injury appears to be a determinant factor foraxonal regeneration within PNGs (1, 5, 53, 54). Forexample, distal axotomy, which elicits minimal peri-karyal reaction with a lower rate of cell death or cellatrophy than that observed following proximal axotomy,appears to entail unfavorable conditions for axonalregeneration.Axons entering PNGs inserted within thebrain or spinal cord tend to originate from nearbyneurons, and regeneration from axons with cell bodiesseveral centimeters from the site of injury has rarelybeen observed (1, 2, 4, 5).The aim of the present work was to investigate the

EXPERIMENTAL NEUROLOGY 137, 1–14 (1996)Article No. 0001

1 0014-4886/96 $12.00Copyright r 1996 by Academic Press, Inc.

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extent to which blind-ended spinal PNGs could stillinduce axonal regeneration of central respiratory neu-rons, even when implanted distant to the respiratorymedullary centers at the level of descending cervicalrespiratory pathways.We have studied axonal regenera-tion within these PNGs inserted into the cervical spinalcord using several techniques. We recorded spontane-ous unitary activities from regenerated fibers withinthe grafts. The CNS origin of these regenerating neu-rons was determined using retrograde labeling withhorseradish peroxidase (HRP). The structural organiza-tion within the PNGs was assessed by light and elec-tron microscopy. Normal patterns of periodic respira-tory discharges were recorded from regenerated axonswithin the grafts. Some of these discharges were provedto be efferent and to originate from central respiratoryneuronswhich remained functional and preserved affer-ent connections. Presence of inflation-related respira-tory discharges suggested that respiratory afferentsalso regenerated within the grafts. Numerous myelin-ated and unmyelinated fibers were observed within thegrafts, and HRP-stained neurons were found in spinalcord and many nuclei of the brain stem includingrespiratory structures. Correlations between electro-physiological and anatomical data are discussed.

MATERIALS AND METHODS

Chronic Graft Implantation

Thirteen female Sprague–Dawley rats weighing 260–300 g (Iffa Credo, Les Oncins, L’Arbresle, France) wereused. The experiments were approved by the nationalauthorities and rats were housed in accordance withFrench law concerning animals. Six rats were used forelectrophysiological investigations and 7 for anatomi-cal studies. In both groups, anesthesia was inducedwith sodium pentobarbital (Nembutal, ip 60 mg/kg),and atropine (ip 1 mg/kg) was administered to reducesecretions. Under aseptic conditions and with the use ofthe dissecting microscope, a hemilaminectomy wasperformed and the meninges were slit longitudinally toexpose the dorsolateral surface of the second segmentof cervical spinal cord (C2). A segment of commonperoneal nerve, approximately 4 cm long was removedfrom the left leg, as previously described (26). One endwas pushed with a fine glass rod into the cervicalenlargement of the spinal cord at the C2 level, on theleft side, and stitched (Ethilon, 10/O) to the overlyingmeninges. Graft implantation was made perpendicularto the dorsal surface of the spinal cord, 1.2–2 mmlateral to the midline and 1–1.5 mm deep; this corre-sponds to the region where medullary respiratoryneurons project their axons within the spinal cord (15,16). The distal part of the graft was left free under theskin and tied with a silk suture (Trinyl, 3/O). Muscles

and skin were closed with 3/O sutures (Trinyl).Animalswere housed in smooth-bottomed plastic cages at 22 61°C with a 12-h light/dark cycle. Food (Purina rat chow)and water were available ad libitum.After a postgrafting period ranging from 2 to 4

months, by which time respiratory neurons are knownto have given off regenerated axons into medullaryPNGs (23, 24, 26, 33), the animals were reanesthetizedwith Nembutal (ip 48 mg/kg) and prepared for electro-physiological examination of the firing pattern, label-ing of neurons sending regenerated axons into thegraft, and graft electron microscopy (Fig. 1A).

Electrophysiology

In this group (n 5 6), a tracheotomy was performedto maintain an unobstructed airway, and the rats wereartificially ventilated with a mixture of room air (50%)and oxygen (50%) (rate 40–60/min, tidal volume 2–4ml). Animals were paralyzed by an iv injection ofgallamine triethiodide (Flaxedil, iv 10 mg/kg). Trachealpressure (T.P) was monitored with a pressure trans-ducer. The right femoral vein was cannulated foradministration of supplemental anesthetic and drugs.Arterial blood pressure was monitored continuouslyfrom the left femoral artery by means of a polyethylenecannula connected to a pressure transducer. Heart ratewas controlled from the arterial pressure pulse. Thelevel of anesthesia was controlled by monitoring bloodpressure, heart rate, pupil size, and reaction to painstimuli. Supplementary doses of anesthetic (Nembutal,iv 4.8 mg/kg) were given at approximately hourlyintervals tomaintain a slow and regular central breath-ing pattern and stable arterial blood pressure (between80 and 120 mm Hg). To monitor the central respiratoryrhythm the left phrenic nerve was exposed by dorsolat-eral approach, cut distally, and placed on tungstenbipolar electrodes. Respiratory rhythm was usually notsynchronized with the pump but ventilation was ad-justed to maintain a pattern of phrenic nerve dischargesimilar to that recorded before paralysis. The chemicaldrive during both spontaneous and artificial ventila-tion was thus similar. Rectal temperature was moni-tored with a probe and kept at about 38°C by a heatingpad. The nerve graft was separated from the surround-ing tissues. After 2–3 mm was removed for light andelectron microscopy, the new end was immersed in apool of warm paraffin oil formed by the surroundingtissues. The nerve graft was desheathed at the distalend. Small strands were teased from the graft formonopolar recording of unitary activity with a silverelectrode referred to a nearby ground. Strands wereteased until the entire graft had been sampled. Phrenicactivity (Phr.N) and axonal activity (Graft Unit) wereamplified (0.5 to 2 K for Phr.N and 2 to 50 K for GraftUnit) and filtered (50 to 3000 Hz for Phr.N and 500 to10,000 Hz for Graft Unit). Phrenic activity was fed into

2 DECHERCHI, LAMMARI-BARREAULT, AND GAUTHIER

a ‘‘leaky’’ integrator (RC integrator with a time con-stant of 50 ms). All the signals [Phr.N, integratedphrenic activity (Int.Phr.N), Graft Unit, T.P] weremonitored on a 4-channel oscilloscope (Gould, DSO1604), chart recorded (Gould, colorwriter 6120 or Astro-med, Dash VIII), and stored on magnetic tape forsubsequent analysis.At the end of each experiment, thespinal cord was removed. After fixation in 10% forma-line, frozen sections (thickness 60 µm) were cut in acoronal plane, stained with cresyl violet, dehydrated,and examined histologically.

Identification and Classification of RecordedGraft Fibers

Graft units were identified as respiratory units (R) ornonrespiratory units (NR) based on a comparison oftheir discharges with that of the phrenic nerve duringartificial ventilation. The inspiratory (I) phase is de-fined as the time during the phrenic burst and theexpiratory (E) phase as the silent period betweenphrenic bursts. Any persistent graft periodic activityobserved in respiratory phases during brief arrest ofthe artificial ventilation and in the absence of anypassive movements by the paralyzed respiratorymuscles was considered to be driven by the centralrespiratory pattern generator and taken to constitutecentral efferent respiratory activity. The activity of Runits therefore reflected the spontaneous discharge of

central efferent respiratory neurons, the axons of whichhad regenerated within the PNGs.The various respiratory discharge patterns were

classified as described previously for central respira-tory neurons (9, 18, 25) and for respiratory neuronsregenerating axonswithinmedullary PNGs (23). Briefly,the phasic patterns of discharge were named all whenthe discharge occurred during the entire I or E phase(generally with either an augmenting or a constantfrequency), late when activity started to fire in mid-phase or later (with an augmenting discharge), earlywhen activity started early during the phase andstopped before the end of the phase with a decrement-ing discharge pattern, and phase-spanning when activ-ity was maximum at the I–E or E–I phase transition.Tonic neurons discharged continuously with a peakfrequency during either I or E phase.Neurons regenerating axons into the grafts were

tested for their responses to modification in pulmonaryfeedback and muscle respiratory afferents by stoppingthe ventilator and/or by inflating (or overinflating) thelungs during E; overinflation prolonged expiration(Breuer–Hering reflex).

Retrograde HRP Labeling of Central NeuronsRegenerating Axons

In the second group (n 5 7), HRP (Sigma type VI)was applied to the extracranial end of grafts to retro-

FIG. 1. (A) General experimental design. The figure shows a blind-ended autologous peripheral nerve graft (PNG) inserted into the left C2spinal cord at the level of the descending respiratory pathways controlling the respiratory motoneurons. After intervals of time to allowneurons from the central nervous system (CNS) to regenerate their axons within the graft, reinnervation of the PNG was investigatedfollowing electrophysiology (a), HRP labeling (b), and electron microscopy (c). For electrophysiological examination, the unitary activity ofregenerated axons was recorded (Graft Unit) after teasing nerve filaments from the graft and compared with the phrenic nerve activity(Phr.N) which was taken as the control of the central respiratory rhythm. (B) Schematic reconstructions illustrating the graft implantation inthe spinal cord at the C2 level in six animals. Drawings were constructed from a number of serial sections through the graft and thus representthe maximum extent of each implant site. In all cases the tip of the grafts reached the left ventrolateral funiculus.

3REGENERATION OF RESPIRATORY PATHWAYS WITHIN PNGs

gradely label cells that extend axons through thegrafts. Each animal was anesthetized (Nembutal, ip 48mg/kg), the skin was reopened, the end of the graft wasexposed, and 2–3 mm was cut off for light microscopy.Tissue surrounding the graft was covered with Para-film sheets and Vaseline to prevent spread of the tracerinto the muscle.AGelfoam pledget soaked in HRP (45%HRP dissolved in 2% dimethyl sulfoxide) was thenapplied to the cut end for 20 min. The pledget wasremoved and the skin wound was closed with sutures.For the control experiments, grafts were exposed andligatured in their midportions and, immediately after-ward, HRPwas applied to the distal end of the grafts.After 72 h, the animals were reanesthetized (Nembu-

tal, ip 60 mg/kg) and sacrificed by intracardial perfu-sion through the aorta with 500 ml of a saline wash (0.1M phosphate buffer, pH 7.2–7.4, 1% procaine HCl, and1% heparin) followed by a mixture of a fixative contain-ing 1.5% paraformaldehyde and 3% glutaraldehyde in500 ml of the same buffered solution at room tempera-ture. The spinal cord and the brain were removed fromC5 to superior colliculus and postfixed 2 h at 4°C in thesame fixative.For routine light microscopy and identification of

retrogradely labeled neurons, the sample was cryopro-tected overnight by immersing it in phosphate buffercontaining 20% sucrose and stocked at 14°C. The nextday it was immersed in isopentane, frozen 15 min at220°C in the cryostat chamber (Microm) and sectionedcoronally. All sections between C3 and pons (a segment18 mm long; 211,150 to 16850 µm from the obex) werereacted with tetramethylbenzidine (TMB) and hydro-gen peroxide (41). Sections 50-µm thick were brieflywashed with distilled water and incubated 20min in anaqueous solution containing 0.1% sodium nitroprus-side, 0.005% TMB, 2.5% ethanol, and 5% acetate buffer,pH 3.3. Hydrogen peroxide (0.1%) was added and thesections were incubated for a further 20min. They werethen mounted on glass slides coated with a chrome-alum gelatin, counterstained lightly with 1% neutralred (TMB procedure), and dehydrated through a gradedseries of alcohol to xylene. Coverslips were applied withthe mounting di-n-butylphtalate xylene (DPX mount-ing media, Fluka Chemical Co., Ronkonkoma, NY). Thesections were examined with a Polyvar microscope.Morphological features of some of these cells wereillustrated by camera lucida drawings or photomicrog-raphy (KodakColor 200/400/600ASA). The labeled nervecells were clearly identified by dark blue peroxidasegranules filling the perikaryal cytoplasm and proximalsegments of dendrites and axons. Special care wastaken to avoid double counting: when the same neuronwas visible in adjacent sections, it was counted onceonly. The distribution of neurons retrogradely labeledthrough grafts was plotted (location of labeled cellbodies was determined from measurements along mid-

sagittal and horizontal coordinates with the center ofthe central canal as a zero reference point), includingthe location of the graft, and data were fed into acomputer. For purposes of description and illustrationof results, the transition between the upper cervicalspinal cord and the beginning of the medulla oblongatawas considered to occur at 21800 µm from the obex ofthe medulla (45).

Electron Microscopy

Samples of nerve graft (2–3 mm cut from the distalstump) were plunged in prefixative (12 h at 4°C in 3%buffered glutaraldehyde), stored (12 h at 4°C in buff-ered sucrose, pH 7.4), osmicated (postfixation for 1 h in2% OsO4), washed in bidistilled water and bufferedsucrose, dehydrated through a graded alcohol series,and embedded in resin mixture (Epon 812 or Araldit).Semithin (1 µm) and ultrathin (60 nm) sections werecut with a glass or diamond knife on an ultramicrotome(Reichter). Sections were mounted on glass blades(semithin) or copper grids (ultrathin). Semithin sec-tions were stained with p-phenylenediamine and ob-served under a light microscope (Polyvar). Ultrathinsections were stained with uranyl acetate (2% in etha-nol) and lead citrate solutions and observed in anelectron microscope (Hitachi Hu-11E). The ultrastruc-ture of the nerve grafts was illustrated by photomicrog-raphy (Tmax Kodak 100 ASA). Myelinated fibers werecounted in 54 standard areas (7 3 8 µm) with standard-ized microscopic fields defined by a morphometric gridapplied on each semithin photograph printed at maxi-mummagnification. On this grid, each field was distrib-uted with a systematic, well-ordered arrangement. Thenumber of myelinated fibers in these areas was used tocalculate the mean density of myelinated fibers/100µm2 in each graft. The area of the grafts was estimatedon micrograph images of complete transverse sectionsphotographed at low magnification. Corrections forshrinkage were taken into account for all calculatedareas. For comparison with the normal nerve, in someanimals peroneal nerves were removed and treatedsimilarly.

Statistics

Data related to recorded activities and HRP-labeledcell and fiber density in the graft were expressed 6SEM. Statistical analysis was performed using a com-mercially available software program (software graph-PAD instat 1.14). Differences were considered signifi-cant at P , 0.05, using the nonparametric unpairedone-tailed or two-tailed P test (Mann–Whitney U two-sample test). The one-tailed test was used when thedirection of the difference of the samples was never indoubt in each case and only answered the question ofthe magnitude of the difference, i.e., whether it was


statistically significant. The one-tailed test mentionedwas indicated just after the P value when this test wasused. The two-tailed test was used when there wasdoubt about the direction of the difference of thesamples.

RESULTS

Animals and Graft Implantation

Grafted animals survived without any apparent neu-rological deficit except for paralysis of the left hindlegdue to the removal of part of the peroneal nerve. Atautopsy, all the grafts were verified to have remainedfirmly and continuously attached to the dorsolateralpart of the cervical spinal cord. The tip of each PNGreached the ventrolateral tracts of the cervical spinalcord (Fig. 1B). The grafts were anatomically integratedonto the host CNS and cysts and cavitation of the spinaltissue were generally not seen in the vicinity of thePNGs.

Electrophysiology

All six spinal PNGs were thoroughly investigatedand contained regenerated axons with propagatingspontaneous action potentials. The mean number offilaments teased from each graft was 93.7 6 17.3. On atotal of 562 teased filaments, 398 (72.7 6 3.4% onaverage) were silent and 164 (27.3 6 3.4%) had unitaryactivity. These unitary discharges were preferentiallyrecorded from filaments located around vascularizedgraft areas. Most (144/164) of the unitary dischargeswere spontaneous during both artificial ventilation andtemporary arrest of the ventilator; discharges wererespiratory (n 5 32) and nonrespiratory (n 5 112). Theother unitary discharges (20/164) resulted from lunginflation (inflation-related units, IR) and consequentlytheir respiratory-related discharge pattern did not

have a central origin. This will be described further inmore detail. The distribution of R, NR, and IR units ispresented in Table 1.Most (23/32) R units were inspiratory; the remainder

were expiratory. All but one of the inspiratory neuronsand all the expiratory presented phasic pattern ofdischarge. The distribution of the various respiratorydischarge patterns is illustrated in Table 2 and ex-amples are shown in Fig. 2. These patterns are similarto those previously described for medullary respiratoryneurons in rat (11, 17, 31, 44, 52) or at the level of uppercervical spinal cord (37). To test peripheral transsynap-tic influences on the activity of central respiratoryneurons regenerating axons within spinal PNGs, wemonitored the effects of lung inflation applied duringthe expiratory phase on the discharge of the R units.Most (29/32) R units were unaffected; they thereforeresemble Ra neurons which receive inputs solely fromthe respiratory central pattern generator (18). Of thethree R units affected by inflation, two (I late) wereexcited. The discharge that normally occurred duringthe last part of the phrenic burst could also be elicitedby lung inflation applied during the E phase (Fig. 2B1).The other unit (an E late) was inhibited (Fig. 2B2).Their cell bodies therefore receive inputs both from thecentral respiratory pattern generator and from respira-tory afferents (probably pulmonary slowly adaptingreceptors). These neurons responded like central neu-rons in the medulla (the so-called Rb) (11, 18, 52) andwithin the spinal cord (10).Most spontaneously active units (112/144) were NR.

They were not simply R units with low levels of activitydue to insufficient respiratory drive, because asphyxia(which increased phrenic activity) did not affect theirdischarge patterns. They discharged phasically, sporadi-cally, or tonically (Table 1 and Fig. 3A). Some tonicunits had discharge patterns synchronized with thecardiac rhythm. For each graft, the number of NR unitswas always higher than the number of R units and themean number of NR units was statistically signifi-

TABLE 1

Unitary Discharges within PNGs

Type of graft unit

Pattern

TotalPhasic Tonic Sporadic

Respiratory (R) 31 1 — 32Nonrespiratory (NR) 27 75 10 112Inflation related (IR) 20 — — 20Total 78 76 10 164

Note. Type and number of unitary activities recorded within spinal‘‘blind-ended’’ peripheral nerve grafts examined between 2 and 4months postgrafting. All these units were recorded in paralyzed andventilated animals and fired with a regular periodic discharge(phasic) or continuously (tonic) or intermittently (sporadic). The Rand NR units continued to discharge during temporary arrest of theventilator.

TABLE 2

Discharge Patterns of Respiratory Units

R unit

Pattern

Total

Phasic

TonicEarly All LatePhase

spanning

Inspiratory (I) 5 13 2 2 1 23Expiratory (E) 2 1 6 0 0 9Total 7 14 8 2 1 32

Note. All these I and E patterns of discharges observed in graftunits corresponded to the activity of central efferent respiratoryneurons that had regenerated axons within the PNGs.


cantly higher than those of R units (P 5 0.032, one-tailed test). The origin(s) of NR activities was notdetermined but similar discharges have also beenreported in PNGs implanted within the brain stem (23,24, 33, 42, 49).

Inflation-related (n 5 20) units fired only in responseto each lung inflation. They were found in all grafts,representing 3 6 0.6% of all recorded filaments. Whenlung inflation and phrenic activity were synchronized,the pattern of activity of these units resembled those ofI units but turning off the ventilator abolished theirdischarge (Fig. 3B1), even under conditions of in-creased respiratory drive (asphyxia). Conversely, theydischarged during maintained lung inflation (Fig. 3B2)

FIG. 2. Examples of respiratory (R) units. All these units pre-sented spontaneous patterns of respiratory activity which persistedwhen the artificial ventilation was temporarily stopped in paralyzedanimals (arrows). These discharge patterns corresponded to theknown pattern for normal central efferent respiratory neurons. (A)Inspiratory (I) and expiratory (E) activities (Graft Unit) with I all(A1) and E late (A2) pattern of discharge. These activities wereunaffected by lung inflation artificially applied during the E phaseand corresponded to the discharge of central respiratory neuronsreceiving inputs solely from the central pattern generator. (B)Examples of R units receiving central and peripheral inputs. In B1,artificial lung inflation applied during the E phase elicits activationof the discharge of the I late unit (asterisks). In B2, the discharge ofthe E all unit is inhibited following lung inflation applied during theE phase (asterisk). Phr.N, phrenic nerve activity; Int.Phr.N, inte-grated phrenic nerve activity; Graft Unit, unit recording; T.P.,tracheal pressure.

FIG. 3. Examples of nonrespiratory (NR) and inflation-related(IR) graft units. (A) Different types of NR units. These unitsdischarged sporadically (A1), phasically (A2), or tonically (A3) evenwhen the ventilator was temporarily stopped. (B) IR unit. In B1, thisunit discharged during each lung inflation but stopped dischargingwhen the artificial ventilation was temporarily halted (arrows). Thisindicates that this discharge pattern is not of central origin. In B2, amaintained lung inflation applied during the E phase elicits along-lasting activation of the unit (asterisk). Abbreviations as inFig. 2.


and their discharge intensity was related to inflationvolume. When the ventilator and phrenic rhythm weredesynchronized their discharge was no longer in phasewith that of the phrenic nerve. The discharge of theseinflation-related fibers resembles that of respiratoryafferents as well as that of central medullary cellsrelaying information from respiratory afferents, i.e.,second-order afferents (11, 52).We tested activity in 93.7 6 17.3 filaments per graft:

66.3 6 11.6 were silent, 18.7 6 4.5 were NR, 5.3 6 1.9were R units, and 3.36 0.8 were IR (Fig. 4A). Figure 4Billustrates the mean occurrence rate of R units refer-enced both to the total number of spontaneous units(R/R1NR; 25.3 6 6.3%) and to the number of teasedfilaments (R/T; 5.9 6 1.6%). Comparisons of R, NR,R1NR, and R/R1NR between spinal grafts and medul-lary ventrolateral (R 5 11 6 3.8; NR 5 14.6 6 7.3;R1NR 5 25.6 6 10.4; R/R1NR 5 57 6 11.4%) or ven-tromedian (4.2 6 1.7; 20 6 4.1; 24.2 6 4.8; 17.7 6 6%)grafts (24, 33) revealed no statistically significantdifferences except for themedullary dorsolateral (16 1;12.8 6 3.8; 13.8 6 3.6; 8.3 6 8.3%) grafts (24) whichcontained a significantly lower mean number of R units(P 5 0.03).

Retrograde HRP Labeling of RegeneratedCNS Neurons

The localization, distribution, and number of CNSneurons that had extended their axons into blind-

ended grafts was determined by studying the retro-gradely labeled cells after application of HRP to theextraspinal portion of the grafts. Labeled cells locatedwithin CNS structures normally containing respiratoryneurons were considered as putative central respira-tory neurons having regenerated their axons within thegrafts.Positive labeling was observed in six of seven ani-

mals. Because no labeled neurons were identified incontrol experiments, we assumed that each HRP-containing neuron had been labeled by axonal retro-grade transport. In those six animals, retrogradelylabeled neurons (total number 1997) were encounteredas far as around 14 mm rostrally and 2 mm caudal tothe graft insertion site. These cells were found bothwithin the spinal cord (n 5 1423) and within the brainstem (n 5 574) with a significant ipsilateral distribu-tion (n 5 1491; P 5 0.046, one-tailed test). Distributionof all labeled cells along the rostrocaudal neuraxis wasbimodal and, both for ipsilaterally and for contralater-ally labeled cells, peaks of distribution were seenaround the spinal insertion site and at the medullarylevel (Fig. 5). Regarding the rostrocaudal distribution,the highest number of neurons was located rostrally tothe graft site (n 5 1519; mean percentage per graft65.5 6 12.3%), whereas labeled neurons seen caudallyto the graft were located in the first two caudal millime-ters (n 5 478; 34.4 6 12.3%). Below this level, no la-beled neurons were seen. Regarding the general distri-bution, the vast majority of the cells were concentratedin the 2 mm surrounding the graft site (n 5 1129;66 6 9.7%). In this area, the number decreases withincreasing distance from the implantation site (Fig.5B). Total HRP-stained (n 5 42) cells in spinal andmedullary structures, where respiratory cells had beendescribed, represented on average 5.8 6 2.5% of alllabeled cells (mean number per graft 7 6 2.3).In the brain stem, labeled cells were confined essen-

tially in the ipsilateral areas (503/574) and a smallernumber of neurons were found to be located contralater-ally (71/574). All neurons were found in structuresknown to project to the spinal cord: the area of thelateral reticulae, the vestibular spinal nucleus, andthat of the brain stem nuclei associated with therespiratory function (nucleus ambiguus and nucleussolitarius). Data relating to distribution of neurons inthe spinal cord and brain stem appear in Table 3. Cellslabeled (n 5 31) in medullary respiratory structuresrepresented on average 4.4 6 2.8% of all labeled cells(mean number per graft 5.2 6 1.8) and were mainlysituated ipsilaterally to the graft implantation both inthe nucleus ambiguus (19/21) and in the nucleus soli-tarius (7/10). Example of heavily labeled neuron withinthe nucleus ambiguus is shown in Fig. 6A.In the spinal cord, labeled cells were identified

dorsally (670/1423) and ventrally (753/1423), i.e., above

FIG. 4. Number and occurrence rate of graft units. In A, thediagrams show the respective mean number per graft of teased (T)and silent (S) filaments compared to the mean number of inflation-related (IR), nonrespiratory (NR), and respiratory (R) graft units. InB, the diagrams show the mean occurrence rate of R units per graft.This occurrence is referenced to the total number of spontaneousunits (R/R1NR) and to the number of teased filaments (R/T).


or below (Fig. 6B) the central canal in the gray matter,with a predominant ipsilateral distribution (n 5 988ipsilaterally and n 5 435 contralaterally). Example oflabeled neurons is shown in Fig. 6B. Cells labeled(n 5 11) in lamina VII, where spinal respiratory cellshad been described in the rat (10), represented in mean1.4 6 0.7% of all labeled cells (mean number per graft1.8 6 0.6) and were exclusively situated ipsilaterally tothe graft implantation.

Anatomical Graft Organization

Grafts were filled with unmyelinated andmyelinatedfibers, Schwann cells, fibroblasts, and blood vesselswith hypertrophied endothelial cells and pericysts.Regenerated fibers were preferentially distributedaround well-vascularized graft areas. The overall ap-pearance of the graft nerve-like structure was similarto that of peripheral regenerating nerves (8) and othertypes of PNGs (7, 30). However, the internal organiza-

tion was strikingly different from that of a normalperoneal nerve. Among the regenerated fibers, unmy-elinated and myelinated axons were distributed inclusters (Fig. 7). As in peripheral nerves, unmyelinatedaxons within the grafts had smaller diameter andmyelinated axons had larger diameter. Heavily myelin-ated large axons frequently found in normal adultperoneal nerve (Fig. 7A) were rarely seen in PNGs (Fig.7B); instead many smaller axons were myelinated.Small axons of similar size in normal peripheral nerveswere unmyelinated or thinly myelinated. Thus, theaxonal density of myelinated fibers was higher withinnerve grafts (4.7 6 0.4/100 µm2) than in a normalperoneal nerve (1.3 6 0.1/100 µm2).At the ultrastructural level, all regenerating axons

contained neurotubules and neurofilaments and werelocated without exception within the basal laminascaffolds. These regenerated axons were always sur-rounded by Schwann cells with which they are associ-ated; no axons growing exclusively on basal lamina orin relation with cells other than Schwann cells wereobserved. The presence of myelin loops, the appearanceboth of large number of perineurial cells and of vastamounts of collagen fibrils normally absent within theendoneurium constitutes typical signs of regenerationwhich appear to be a recapitulation of primarymyelina-tion in the developing nerve (32). Characteristic pro-files of regenerating neurites and growth cones withnumerous microtubules, neurofilaments, vesicular pro-files, and mitochondria were not observed.

DISCUSSION

These experiments constitute the first demonstra-tion that PNGs inserted into the cervical (C2) spinalcord can promote regeneration of efferent and afferentrespiratory pathways. The efferent respiratory neuronsregenerating axons retained their original function andcontinued to fire spontaneously during each respiratorycycle with discharge patterns similar to those of normal

FIG. 5. Distribution of horseradish peroxidase (HRP)-labeledneurons in the CNS. (A) The drawing shows a computer-assistedserial reconstruction of a dorsal view of the CNS (brain stem andupper cervical spinal cord) in which each dot represents a labeledneuron which has regenerated an axon into the nerve grafts (n 5 6).(B) The histograms show the distribution of labeled cells along thecaudorostral axis in the ipsilateral and contralateral side. Note thelow number of labeled cells at the level of the graft site (asterisk).

TABLE 3

Distribution of HRP-Labeled Cells

Spinal cord Brain stem Total

Ipsi- 164.6 6 86.3 83.8 6 49.3 248.5 6 123(54.9 6 6.6) (21.2 6 8.2) (76.1 6 6.9)

Contra- 72.5 6 66.1 11.8 6 7.2 84.3 6 73.1(17.8 6 7.5) (6 6 3.8) (23.9 6 6.9)

Total 237.2 6 149.3 95.7 6 53.6 332.8 6 187(72.7 6 9.5) (27.3 6 9.5)

Note. Spinal and brain stem central neurons labeled after applica-tion of HRP to the distal cut end of the cervical PNGs (n 5 6). Labeledcells were located ipsilaterally (Ipsi-) and contralaterally (Contra-) tothe graft insertion site. Values are mean numbers of labeled cells 6SEM. In brackets, percentage of labeled cells.


FIG

.6.

Examples

ofhorseradish

peroxidase

(HRP)-labeledneuronsat

themedullaryandspinal

level.Thepreparationswere

counterstained

withneutral

redandphotom

icrographsof

sectionsweretakenin

dark

field.

HRP-labeled

neuronsappear

loaded

with

chromogen

granulesfillingtheperikaryalcytoplasmas

wellasproximalsegm

entsofdendrites

andaxons.(A)L

abeled

cellatthelevelofthe

nucleusam

biguusas

indicatedinthedraw

ing.Thiscellisconsideredas

aputative

respiratoryneuron.(B)Spinalcellslabeledcontralaterally

tothesiteofgrafting.Scalebar,30

µm.Abbreviations:AP,area

postrema;Sol,nucleusofthesolitary

tract;Amb,am

biguusnucleus.

FIG. 7. Histology of normal and grafted nerve. (A) Normal peroneal nerve. (B) Nerve graft examined 3 months posttransplantation.Semithin (1) and ultrathin (2) sections. Heavy myelinated fibers with large diameter are regularly arranged in peroneal nerve and nuclearprofiles are unfrequently observed. In the nerve graft, note the presence of numerous regenerated myelinated fibers with small size compared

10

respiratory neurons. In addition, their responses dem-onstrated continued inputs from peripheral and centralsource, a result identical to that using PNGs insertedinto the medullary respiratory centers (23, 24, 33, 34).Regenerating respiratory afferents, although not stud-ied in detail, had discharge patterns similar to thosedescribed in normal conditions (14). HRP labelingconfirmed the central origin of neurons extending axonsinto the PNGs and light and electron microscopy re-vealed numerous unmyelinated and myelinated fiberswithin the grafts. The large quantity of small myelin-ated axons within the grafts was reminiscent of whatoccurs during nervematuration and regeneration, whenthe increase in size of the myelinated fibers is precededby an increase in the number of small myelinated fibers(8, 27). However, the absence of regenerating profiles atthe time of examination (2–4months postimplantation)suggested that all axons had already reached the distalpart of the grafts and were not in the growth phase.The discussion will focus on the following: (1) neuro-

nal types regenerating fibers within the grafts, (2)features of axonal regeneration specific to central respi-ratory neurons, and (3) general aspects of CNS axonalregeneration based on both electrophysiological andanatomical approaches.

Neuronal Types Regenerating Fibers within the Grafts:General Features

Different types of activity were recorded from regen-erated axons within the grafts (NR, IR, and R); conse-quently, various neuronal populations were involved inaxonal regeneration but only IR and R fibers could beidentified unambiguously based on their distinctivedischarge patterns.Most (73%) of the filaments isolated from the grafts

were silent. There are two possible explanations for theinactivity; either the neurons would be active onlyduring other behaviors (e.g., locomotion) or the func-tional state had changed. Indeed, changes in mem-brane properties and alterations and rearrangment ofsynaptic inputs are induced by axotomy (35, 36, 40) orinjury to afferents (49) produced by the grafting proce-dure (21, 50).NR fibers originated from neurons which conserved

at least some functional activity since they were stillable to generate spontaneous action potentials. Sincetheir function could not be established under ourexperimental conditions, we do not know if thesespontaneous discharge patterns were ‘‘normal’’. How-ever, some had discharge patterns (e.g., synchronizedwith the cardiac rhythm or rhythmic bursting), suggest-

ing that their cell bodies had conserved part of theiroriginal function.IR fibers discharged during lung inflation, the dis-

charge intensity being related to inflation volume.These fibers discharged like medullary second orderafferents (Pump, or P, cells) relaying volume-relatedvagal feedback. Since these neurons are located exclu-sively in the medulla oblongata and do not project tothe spinal cord (11, 52), we think that IR fibers origi-nate from regenerating respiratory afferents. The sourcecould be either from vagal afferents projecting directlyfrom the nodose ganglion to the upper cervical spinalcord in the rat (22, 39) or from afferents of the dia-phragm and intercostal muscles (13, 14). Identificationof putative vagal afferences following bilateral va-gotomy was not performed in order to maintain experi-mental conditions similar to those in which medullarygrafts were analyzed (24, 33). Unfortunately, reversibleblocking of the vagal information by cooling the vagusnerves could not be carried out for technical reasons.Although the ability of peripheral fibers to regenerateinto spinal nerve grafts has previously been establishedthrough anatomical studies (19), our study provides thefirst evidence that some afferent fibers regeneratingwithin PNGs retain normal functional properties.

Medullary and Spinal Origin of Regenerating CentralRespiratory Neurons

Respiratory neurons regenerating axons within spi-nal PNGs were axotomized at the C2 level (the site ofinsertion of the graft). Presumably, therefore, the graftwas inserted into the descending respiratory pathwaystransmitting central respiratory drive to respiratorymotoneurons. Two main descending spinal respiratorypathways controlling the spinal respiratory phrenicand intercostal motoneurons have been described inthe rat. The first originates from brain stem (pons butmainlymedulla oblongata) neurons projectingmonosyn-aptically to phrenic motoneurons (12, 15, 16, 47). Theother pathway originates from other neurons (cortex,pons, medulla oblongata) which relay on segmentalinterneurons in the upper cervical (C1–C3) spinal cord(3, 10, 37), bypass the phrenic nucleus, and project tointercostal motor nuclei (37).Several of our findings, particularly the location of

HRP-labeled cells in both the medulla and the uppercervical spinal cord, suggest that respiratory neuronsregenerating axons within the PNGs originate fromthese two locations. Thus, R units have dischargesrepresentative of the activity in both descending path-ways.Although we cannot be certain, HRP-labeled cells

to the normal peroneal nerve. These regenerated fibers are clustered, numerous myelin loops are formed around axons, and Schwann cellnuclear profiles are frequently observed. Nerves were examined at the level of their distal part. Scale bars, 12 µm for semithin sections (1) and4 µm for ultrathin sections (2).


inmedullary respiratory areaswere likely those project-ing to the spinal cord, particularly the phrenic nucleus.These labeled cells constitute strong evidence that atleast some of the regenerating respiratory fibers withinthe grafts originate from medullary respiratory neu-rons. However, had they been the only source of respira-tory input into the graft, we would have expected lessrespiratory reinnervation within PNGs inserted intothe cervical spinal cord than in grafts inserted into themedulla (24, 33) near regions containing respiratoryneurons (12, 31) because regeneration is less vigorouswhen the graft site is farther away from the cell body (1,21, 50). In fact, there was no significant differencebetween the numbers of R units in spinal and medul-lary grafts, suggesting that spinal respiratory neuronswere probably involved in the reinnervation of PNGsinserted into the spinal cord. Among labeled cells in thespinal cord, those which were concentrated in theC1–C2 lateral border of lamina VII, an area containingupper cervical respiratory neurons (10), could be seg-mental respiratory interneurons which relay centralrespiratory drive to the thoracic spinal cord (3, 10, 37).Although most graft units could not be related to

either respiratory medullary (11, 17, 31, 44, 52) orspinal neurons (37) which present a lot of commondischarge patterns, some graft units may be specificallyattributed to one or another group of regeneratingrespiratory neurons. For example, E units likely reflectthe activity of medullary neurons because spinal cervi-cal neurons with expiratory discharge patterns havenot been described. Conversely, I/E units may have aspinal origin because medullary neurons with thisdischarge pattern and projections in the spinal cordhave not been reported (44).

Anatomical and Functional Correlation on CNSAxonal Regeneration

HRP labeling confirmed the ability of neurons in themedulla and spinal cord to regenerate axons into PNGsimplanted in the rat spinal cord.As in other studies, themajority of regenerating cells appeared to be locatedaround the implantation site (1, 2, 19, 30), indicatingthat growth-favoring conditions in the PNGs mainlyinteract with nearby cells. However, the density oflabeled neurons did not decrease with increasing dis-tance from the site of the graft, since a bimodaldistribution of labeled cells was found in the spinal cordand medulla. The presence of significant numbers oflabeled cells in the medulla can be attributed to theplacement of the graft in the ventral part of the spinalcord and the associated damage to descending path-ways. This trauma, likely greater than that associatedwith implants in the dorsal part of the spinal cord, mayaccount for more labeling of neurons (approximately330 versus 6–200 in other studies) (1, 19, 20, 30).Moreover, we found labeled cells as far as 14mm rostral

to the site of implantation of the PNG, i.e., at distancesobtained only when axonal regeneration was enhancedusing growth factors (19) or injury combined withgrafting (50, 51).Because we used several different approaches, some

correlations between electrophysiological data, anatomi-cal features of the graft, and retrograde labeling dataare possible. First, fibers with respiratory dischargeswere obtained preferentially from vascularized nervousfilaments (or filaments situated near vessels); anatomi-cal analysis revealed preferential regeneration of fibersnear vascularized areas. These findings suggest thatvascularization is important for axonal regeneration.The outgrowth of neurites and the activities of the cellpopulations supporting regeneration are closely relatedto the vasculature supplying nutrients, oxygen, andother necessary factors (46). Second, the percentage ofputative labeled respiratory cells regenerating axons(5.8 6 2.5% of all labeled cells in the medulla andspinal cord) was similar to the percentage of fibers(5.9 6 1.6% of all teased fibers) with respiratory activ-ity. This would suggest that most, if not all, respiratoryneurons retained their functional properties after axo-nal regeneration. Interestingly, the number of CNSHRP-labeled cells is less than the estimated number ofmyelinated fibres in the grafts (approximately 330 pergraft versus more than 4000 fibers calculated on thebasis of an estimated cross-sectional area of the graft).Interpretation of this feature is difficult since thenumber of regenerated fibers within the grafts does notnecessarily reflect the number of regenerating neurons:some fibers within the grafts could have originatedeither from axon collaterals or from supernumeraryaxons of a single neuron (28, 38). However, electrophysi-ological experiments could underestimate the numberof regenerating axons due to damage, and subsequentloss, of spontaneously active axons during their separa-tion from the graft. Similarly, we may also have under-estimated the number of HRP-labeled cells projectingto the graft due to failure to label neurons (43) andbecause cells outside the CNS (e.g., dorsal root ganglioncells) may have been a source of regenerated axons ofperipheral origin (19).

Conclusion

PNGs implanted in the second segment of the cervi-cal spinal cord can promote axonal regeneration ofrespiratory neurons conveying functional efferent andafferent signals. Regenerating axons carrying efferentsignals originated from central respiratory neurons inboth the upper cervical spinal cord and themedulla. Wesuggest that efferent respiratory information repre-sented two descending respiratory pathways: the bulbo-spinal pathway (from the respiratory centers of themedullamainly to phrenicmotoneurons) and the bulbo-cervicospinal pathway (from the respiratory centers of


the medulla to cervical respiratory interneurons andfrom there to the thoracic spinal cord). The presence offunctional respiratory afferents in the grafts supportsour previous hypothesis (26) that PNGs could provide‘‘real’’ functional supplementary nerves with afferentand efferent components. This feature may contributeto efforts to use PNGs, in combination with otherstrategies, for CNS reconstruction.

ACKNOWLEDGMENTS

The authors gratefully acknowledge the helpful advice and criticalcomments on the manuscript from Dr. S. Iscoe. Expert technicalassistance was provided by P. Rega, A. M. Lajard, J. Pio, and M.Manneville. We also thank Professor J. F. Pellissier and E. Pasqualeof the Service d’Anatomie Pathologique et de Neuropathologie de laFaculte de Medecine de Marseille, for their excellent help in electronmicroscopy. This work was supported by CNRS (URA 1832), DRET(Grant 92-053), and IRME (Institut pour la Recherche sur la moelleEpiniere).

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Regeneration of Respiratory Pathways within Spinal Peripheral Nerve Grafts