If ‘generalist’ RVLM neurons exist, it might be predicted that their descending axons branch to innervate preganglionic neurons of a range of functional classes at multiple spinal segments. The present study aimed to investigate this issue, anatomically and functionally, with contemporary tools. We asked the following questions: (i) to what extent do the axons of individual RVLM neurons branch to innervate preganglionic neurons in disparate spinal levels (upper thoracic vs. lower thoracic/lumbar); (ii) do RVLM neurons that project to lower thoracic/lumbar levels also send collateral connections that excite upper thoracic sympathetic outflows; and, if so, do RVLM neurons that innervate both upper thoracic and lower thoracic/lumbar levels represent a subpopulation of neurons that innervate functionally similar but anatomically distant targets, as suggested by McAllen and Dampney ( 1990 ), or do they provide divergent drive to several functional classes of sympathetic outflow?

The two lines of evidence noted above lead naturally to the hypothesis that individual RVLM neurons are functionally dedicated to drive a single class of peripheral sympathetic neuron (e.g. muscle vasoconstrictor or cardioaccelerator). Against this notion is the general observation that RVLM nanoinjections nearly always coactivate more than one peripheral sympathetic neuron class (Beluli & Weaver, 1991b ). Although this finding might be explained by the spatial spread of nanoinjected substances, no experiment has yet excluded the alternative possibility that there are also ‘generalist’ RVLM neurons with descending connections that synapse with more than one functional class of sympathetic outflow. The discovery of such neurons would suggest that an earlier view of the sympathetic nervous system, specifically that of a monolithic and global effector system that ‘seem[s] devised for widespread diffusion of nervous impulses’, should be reconsidered (Cannon, 1915 ).

In addition to the above evidence for central topography based on function, individual RVLM neurons have also been found to display heterogeneous functional properties. For example, there are differences in their sensitivities to stimulation of arterial baroreceptors or somatic nerves (McMullan et al . 2008 ; Verberne & Sartor, 2010 ) and to central respiratory drive (McAllen, 1987 ; Miyawaki et al . 1995 ; Moraes et al . 2013 ). Corresponding diversity is found among different peripheral sympathetic nerve classes (Janig & McLachlan, 1992 ) and, indeed, in the properties of the sympathetic preganglionic neurons (Morrison & Cao, 2000 ; Stalbovskiy et al . 2014 ). One particularly clear example is the sympathetic control of adrenaline secretion, which appears to be controlled by a group of atypical RVLM bulbospinal neurons that are insensitive to baroreceptor stimulation but are activated by glucoprivation (Verberne & Sartor, 2010 ). These cells may selectively drive adrenal nerve activity, and therefore adrenaline secretion, in response to hypoglycaemic stimuli (Korim et al . 2014 ; Korim et al . 2016 ).

Although the cardiovascular actions of RVLM premotor neurons are not in doubt, there is evidence of functional heterogeneity within this population. In the cat, localized nanoinjections of excitatory amino acids into different subregions of the RVLM were found to preferentially or exclusively activate vasomotor sympathetic nerves to different tissues (Dampney & McAllen, 1988 ; McAllen & Dampney, 1990 ; McAllen & May, 1994 ). McAllen and Dampney ( 1990 ) found that vasoconstrictor nerves to forelimb and hindlimb muscles were coactivated independently from renal sympathetic nerves, and concluded that RVLM subregions were probably organized topographically by the function, rather than the anatomical location, of their peripheral vascular targets. Although the limited spatial resolution of the method makes the functional separation of these outputs more difficult in the rat, it has been demonstrated that sympathetic outflows of different functional classes are activated to varying degrees by RVLM nanoinjections in this species as well (Beluli & Weaver, 1991a , b ; Mueller et al . 2011 ).

The rostral ventrolateral medulla (RVLM) contains spinally projecting neurons that monosynaptically innervate sympathetic preganglionic neurons and are a major supply of the essential tonic drive that maintains arterial blood pressure (reviewed by Dampney, 1994 ; Guyenet, 2006 ). When activated experimentally, these neurons raise blood pressure and excite sympathetic nerves innvervating a range of organs and tissues (Dampney & McAllen, 1988 ; McAllen & May, 1994 ; Abbott et al . 2009 ), although they do not appear to drive ‘non‐cardiovascular’ sympathetic classes, such as sudomotor, pilomotor, pupillodilator, brown adipose thermogenic or intestinal motility inhibitory neurons (McAllen, 1986 ; Cao et al . 2010 ).

Stimulus‐triggered averages of rectified and smoothed (time constant = 10 ms) nerve activity were used to identify activation of post‐ganglionic sympathetic nerves. At least 750 sweeps were averaged in each case. Time‐locked peaks in sympathetic nerve activity (SNA) that exceeded the pre‐stimulus amplitude by at least 2 SDs were interpreted as proof of optogenetic stimulation. Negative controls were obtained by repositioning the fibreoptic probe 3 mm dorsal to the RVLM and repeating the same procedure or by disconnecting it from the laser. The pre‐stimulus mean and standard deviation (σ) were calculated using a period of 200 ms before laser stimulation. At the conclusion of electrophysiology recordings animals were perfusion‐fixed. Brainstems were prepared for histology as described above and RVLM hChR2‐mCherry expression was verified.

Only neurons that lay within the anatomical boundaries of the RVLM were included in analysis. The RVLM was defined using a 3D density map of TH‐immunoreactive neurons that were retrogradely labelled with GFP or mCherry ( n = 273); computed at a 1 voxel (39 μm) resolution using a parametric density mapping method (Burguet et al . 2011 ; Burguet & Andrey, 2014 ) and with the smoothing parameter k set to 10. The boundary of the RVLM was defined as the isodensity surface that contained 85% of the TH+ bulbospinal neurons. The RVLM boundary iso‐surface and the Cartesian co‐ordinates of identified bulbospinal neurons were then imported into Imaris, version 8.1 (Bitplane AG, Zurich, Switzerland). Neurons falling within the RVLM were identified using an Imaris script (‘Spots split into surface objects’; http://open.bitplane.com/tabid/235/Default.aspx?id=19 ) and tabulated, allowing elimination of neurons that fell outside of the RVLM. The x , y and z co‐ordinates from each replicate were plotted as the mean ± SEM, grouped and compared in Prism, version 7.02 (GraphPad Software Inc., San Diego, CA, USA) using one‐way ANOVA followed, if significant, by Tukey's post hoc tests between groups. P < 0.05 was statistically considered significant. The 3D spatial distributions of GFP‐, mCherry‐ and double‐labelled RVLM neurons were pooled over replicates and were assessed by extracting the 85% isodensity surfaces from the three corresponding density maps as described above. The proportion of bulbospinal neurons that expressed TH immunoreactivity was compared using a chi‐squared test.

Labelling exhibited some variability between animals: experiments in which retrograde labelling from T 2 or T 10 was conspicuously absent were assumed to reflect off‐target vector injections and were eliminated from analysis. Previous experiments using HSV vectors indicated that retrograde transduction of spinally projecting RVLM neurons was largely restricted to the ipsilateral side (Dempsey et al . 2017 ) and therefore labelling on one side of the brainstem was considered independent of the labelling on the other. Accordingly, although data were collected from both right and left sides of the medulla, data were transformed and pooled such that all were plotted on the left side.

RVLM sections (50 μm) were aligned to the Waxholm volumetric rat brain atlas (Papp et al . 2014 ) using the QuickNII affine registration tool ( https://www.nitrc.org/projects/quicknii ). Labelled neurons lying in a triangle defined by the ventral midline surface, nucleus ambiguus and medial border of the spinal trigeminal tract were manually annotated and classified based on the presence of GFP, mCherry and/or TH immunoreactivity. The pixel co‐ordinates of annotated neurons were transformed into Waxholm co‐ordinates as described previously (Dempsey et al . 2017 ).

Forelimb MVC nerve activity was recoded from the triceps branch of the ulnar nerve. The nerve was dissected from the triceps, cut distally, desheathed and placed over a small black plastic dissection platform under mineral oil. Filaments were split from its central end using watchmaker's forceps and a razor shard. Activity was recorded differentially between an active filament and a strand of connective tissue led over two fine silver wire electrodes. Hindlimb MVC activity was recorded from the gastrocnemius branch of the popliteal nerve using a similar approach. MVC spike activity was identified by its irregular ongoing activity, usually modulated by the cardiac cycle.

The lumbar sympathetic chain on the left side was exposed via a retroperitoneal approach, as described previously (Korim et al . 2011 ), then blunt‐dissected and cut rostral to the L 3 ganglion. The left kidney was exposed and gently retracted. The lumbar sympathetic chain was blunt‐dissected, isolated and cut caudal to the L3 sympathetic ganglion. Its rostral end was placed across two silver wire electrodes and its activity recorded as described above.

The left cardiac sympathetic nerve was isolated as described previously (Pracejus et al . 2015 ). In brief, the head of the second rib was identified and cleared, taking care not to puncture the pleura. The head of the second (and often also the third) rib was removed and the peripheral portion retracted downwards: the stellate ganglion was accessed immediately underneath and was identified by its location and distinctive morphology (Korzina et al . 2011 ). The cardiac sympathetic nerve was identified by its trajectory from the lateral and/or caudal aspect of the stellate ganglion towards the heart and blunt‐dissected free with fine forceps. It was then cut distally as it entered the pleura. It was led over a pair of fine silver wire electrodes under mineral oil and its activity was amplified and filtered (10–100 Hz high pass, 600–1000 Hz low pass; Neurolog, Welwyn Garden City, U.K.) and digitized at 5 kHz using a CED Power 1401 interface and Spike2 software (Cambridge Electronic Design, Cambridge, UK). An earth electrode was placed at the origin of the nerve from the stellate ganglion to minimize ECG contamination.

Eleven to 50 days after vector injections, surgical anaesthesia was induced as described above. Animals were tracheostomized and artificially ventilated with 2% isoflurane in 100% oxygen. The femoral artery and vein were cannulated for the monitoring of blood pressure and i.v. access, respectively, and rats placed in a stereotaxic frame. Paired recordings were undertaken in several combinations: forelimb and hindlimb muscle vasoconstrictor (MVC); cardiac sympathetic nerve and lumbar sympathetic chain; cardiac sympathetic nerve and hindlimb MVC.

Exposed tissue was irrigated with sterile saline, a piece of sterile Gelfoam (Pfizer Inc.) was placed over the exposed surface of the spinal cord, and incisions were closed in layers using silk thread. Anaesthesia was discontinued and animals closely attended until ambulatory. They were then monitored daily signs of spinal cord injury, pain or weight loss.

This intersectional approach enabled us to target those neurons with cell bodies in the region of the RVLM for which the axons project to the thoracolumbar spinal cord: only neurons that were exposed to viral vector at both the central and the spinal injection site would express Channelrhodopsin2 (ChR2).

Animals were then secured in a stereotaxic frame tilted 10° nose‐down and the dorsal aspect of the skull was exposed. A small craniotomy was drilled overlying the injection sites and a micropipette filled with AAV‐EF1a‐DIO‐hChR2‐mCherry (AAV serotype 2), supplied at 1 × 10¹³ vg mL –1 ; Penn Vector Core, University of Pennsylvania, Philadelphia, PA, USA) was advanced into the region of the left RVLM (anteroposterior: 11.96; mediolateral: –2.1; dorsoventral: –10.5). Nanoinjection (100–200 nL) was frequently associated with respiratory artefacts (tachypnoea or bradypnoea), indicating that the injections were in the region of the ventrolateral medulla. After the initial injection, the micropipette was advanced rostrally at intervals of 0.2–0.3 mm until nanoinjection produced twitching of the whiskers, indicating proximity to the facial nucleus, an anatomical landmark for the rostral margin of the RVLM. Once three or four injections had been performed, injections were repeated 0.3 mm more medially at the same anteroposterior co‐ordinates. In total, six to eight injections of 100–200 nL were made spanning the region: mediolateral 1.8–2.1; anteroposterior: 11.3–12.7 mm.

The caudal‐most rib was used as a landmark to identify the T 13 vertebral segment. Overlying muscle was cleared from the T 12 ‐L 1 vertebrae: the spinal level at which sympathetic preganglionic neurons that target hindlimb muscle vasoconstrictor neurons are primarily found (Sonnenschein & Weissman, 1978 ; Baron et al . 1988 ). The T 12 vertebra was clamped for mechanical stability, and a laminectomy was performed. A drop of lidocaine was placed on the spinal cord surface to prevent spinal locomotor reflexes and 250–300 nL of a canine adenoviral vector (CAV2‐CMV‐Cre, supplied at 2.5 × 10 12 pp mL –1 ; Eric Kramer, Plateforme de Vectorologie de Montpellier, Montpellier, France) was pressure injected into the left IML via a bevelled glass micropipette (diameter 20 μm) positioned just lateral to the posterior spinal vasculature and 0.5 mm ventral to the spinal surface. The injection volume was determined by visualization of the fluid level within the micropipette using a microscope fitted with a calibrated graticule. The micropipette was left in place for at least 2 min; six to nine such injections were made per animal at intervals of ∼0.5 mm in the rostrocaudal direction.

In electrophysiology experiments, we examined whether photoactivation of RVLM neurons that project to the lower spinal cord also produced responses in functionally similar and functionally distinct sympathetic nerves that emerge from the upper cord. A schematic diagram presenting our overall hypothesis and experimental strategy is provided in Fig. 4 . Functional studies were carried out in 44 male Sprague–Dawley rats (200–370 g) bred on‐site at the Florey Institute. Animals were under a 12:12 h light/dark cycle and had free access to food and water. Anaesthesia was induced with sodium pentobarbitone (60 mg/kg, i.p.) and maintained with 2–3% isoflurane in oxygen; the criterion for successful anaesthesia was areflexia to a strong pinch. Animals were treated with meloxicam (Ilium; Troy Pharmaceutical, Smithfield NSW, Australia; 0.5 mg, i.m.) and atropine (Pfizer Inc.; 60 μg, i.m.) for analgesia and prevention of excessive tracheal secretion, respectively, then transferred to a homeostatic heating blanket controlled via a rectal temperature probe and maintained at 37 ± 1°C.

In two cases rats, were perfused with modified acrylamide‐paraformaldehyde fixative solution and cleared using the CLARITY protocol (Tomer et al . 2014 ). Forty‐eight hours after perfusion, the fixative was polymerized and the brainstems cut into 1 mm thick coronal sections using a brain matrix. Sections were then incubated with SDS/borate buffer and passively cleared over several weeks before mounting in a custom imaging chamber filled with glycerol. CLARITY‐processed brains were tile‐imaged using a TCS SP5X confocal microscope (Leica Microsystems, Wetzlar, Germany).

At the conclusion of experiments, rats were killed with sodium pentobarbitone (>150 mg kg ‐1 ; Lethabarb; Virbac, Milperra, NSW, Australia) and transcardially perfused with 300 mL of cold heparinized saline followed by 300 mL of 4% paraformaldehyde. The brain and thoracic spinal cord were removed, post‐fixed overnight and the medulla blocked and cut into 50 μm coronal sections in a 1:4 series. For anatomy experiments, sections were permeabilized in 0.2% Triton‐100 (3 × 15 mins) then incubated with primary antibody [T1299 monoclonal mouse anti‐tyrosine hydroxylase (TH); AB_477560; dilution 1:1000; Sigma, St Louis, MO, USA;) for 48 hours in TPBS containing 2% BSA. Sections were then washed (3 × 30 min) and incubated overnight in blocking buffer containing Alexa Fluor® 647‐conjugated secondary antibody (A31571 polyclonal goat anti‐mouse; AB_162542; dilution 1:500; Life Technologies, Grand Island, NY, USA). Sections were washed again in TPBS (3 × 15 min), mounted in order and coverslipped with Dako mounting medium. No immunohistochemical processing was required to visualize GFP‐ or mCherry fluorescence. Brain sections that spanned the RVLM were imaged under epifluorescence (AxioImager Z2 microscope; Carl Zeiss, Oberkochen, Germany; 10×/0.30 NA M27 objective; 0.645 μm pixel size).

After injections the spinal cord was irrigated with sterile saline and covered with Gelfoam (Pfizer Inc., New York, NY, USA). Skin incisions were closed with metal clips and anaesthesia reversed with atipamazole (1 mg/kg s.c.; Pfizer Animal Health) and rats monitored closely until ambulatory. Rats were housed singly for 4–5 days after vector injections with frequent monitoring including daily weighing and behavioural assessment and additional analgesia as required.

In subsequent experiments, HSV‐hCMV‐GFP was used to label T 2 ‐projecting neurons and HSV‐hCMV‐mCherry for T 10 ‐projecting neurons. Two 500 nL injections of each vector were made bilaterally (i.e. a total of 2 μL of each vector). Injections were made over 5–10 min and pipettes left in place a further 5 min to minimize vector leak from the parenchyma.

We first established that HSV‐hCMV‐GFP and HSV‐hCMV‐mCherry vectors result in equivalent retrograde transduction of medullary neurons. This was achieved by injecting a 1:1 mixture of HSV‐hCMV‐GFP and HSV‐hCMV‐mCherry at the T 2 IML (2 × 500 nL injections on each side of the spinal cord separated by 1 mm rostrocaudal) and examining the proportion of medullary neurons that expressed both reporters four days later ( n = 2 animals).

On the day of surgery, animals were anaesthetized with ketamine and medetomidine (75 mg kg –1 i.p.; Parnell Laboratories, Mascot, NSW, Australia; and 0.75 mg kg –1 i.p.; Pfizer Animal Health, Sydney, NSW, Australia, respectively), treated with prophylactic analgesia (carprofen, 2.5 – 10 mg kg –1 s.c.; Norbrook Pharmaceuticals, Tullamarine, VIC, Australia) and antibiotics (cephazolin, 100 mg kg –1 i.m.; Mayne Pharma, Melbourne, VIC, Australia), the fur on the back clipped and the skin scrubbed with betadine. Anaesthesia was assessed by examining withdrawal of the hind paw in response to noxious pinch and was supplemented with ketamine (7.5 mg kg –1 i.p.) as required. Rats were positioned in a stereotaxic frame on a thermostatically controlled heated blanket and the T 2 and T 10 spinal cord exposed by blunt dissection of the overlying muscles and separation of the spinous ligament (T 2 ) or laminectomy (T 10 ). Adjacent vertebrae were clamped for stability. Bilateral pressure injections of HSV‐hCMV‐GFP or HSV‐hCMV‐mCherry vectors (supplied at 3 × 10 8 IU mL –1 ; Rachael Neve, McGovern Institute for Brain Research at MIT, Cambridge, MA, USA) were made at the interomediolateral column (IML) using a glass pipette mounted on a manipulator. Injections were targeted 1 mm deep to the dorsal root entry, ∼0.7 mm lateral to midline. Appropriate co‐ordinates were verified in initial experiments by addition of blue polystyrene beads to the injectate (#09980508; dilution 1:10,000; Thermo Scientific, Scoresby, VIC, Australia).

Evidence for RVLM bulbospinal neurons that innervate multiple spinal segments was sought in experiments in which two replication‐deficient retrograde viral vectors that expressed different fluorophores were each injected into the upper or lower thoracic spinal cord. Adult Sprague–Dawley rats (Animal Resource Centre, Perth, WA, Australia) were housed under 12:12 h light/dark cycles in individually ventilated cages with access to food and water available ad libitum . The data presented in the present study were obtained from 10 male rats (222–380 g), with data rejected from a further six animals as a result of poor labelling. A further 13 male and female rats were used to optimize time course and vector targeting.

As reported by Abbott et al . ( 2009 ), laser‐evoked SNA responses were sometimes followed by post‐burst inhibition that lasted 100–200 ms; periods of reduced nerve activity that exceeded σ by four‐ to six‐fold were apparent in forelimb MVC (two of four animals), cardiac SN (two of eight animals), lumbar SN (one of four animals) and hindlimb MVC (one of six animals). In addition, two recordings of hindlimb MVC showed laser‐evoked inhibition of SNA that was not preceded by activation (5–6σ, latencies of 253 and 275 ms).

We then investigated whether evidence of functional collateralization of RVLM neurons is observable in non‐MVC sympathetic outflows. For this, we again used injection of CAV‐cre in the lower spinal cord combined with injection of AAV‐DIO‐ChR2 in the medulla to control ChR2 expression in the RVLM. We recorded activity from the cardiac sympathetic nerve at the same time as either the hindlimb MVC ( n = 4) (Fig. 6 A ) or L2 lumbar sympathetic chain ( n = 4) (Fig. 6 B ). Intermittent optogenetic stimulation of the RVLM produced pulse‐locked activation of the cardiac sympathetic nerve with amplitudes exceeding those observed during the pre‐stimulus period by 5σ in all eight animals tested. Peak cardiac sympathetic nerve response latencies fell between 53 ms and 158 ms. A double peak (63 and 112 ms) was seen in one animal (Fig. 6 C ). Stimulus‐locked activation of hindlimb MVC was observed in three of four experiments (peak latencies: 180–313 ms), one of which featured a double peak (latencies: 203 ms and 313 ms) (Fig. 6 D ). On average, peaks in hindlimb MVC activity occurred 121 ± 10.4 ms after those of the cardiac nerve. Stimulus‐locked activation of the lumbar sympathetic outflow was observed in four of four experiments (peak latencies: 152–229 ms). On average, peaks in lumbar sympathetic activity occurred 76.5 ± 4.6 ms after those of the cardiac nerve.

Examples of event‐triggered averages of sympathetic nerve activity evoked by laser‐stimulation of the RVLM region (473 nm, 10 mW, 20 ms pulse width, 1–2 Hz). Dotted lines indicate upper and lower control limits., averages of light‐evoked cardiac and hindlimb SNA (control limits: ± 6σ; 2493 sweeps averaged)., averages of light‐evoked cardiac and lumbar SNA (control limits: ± 6σ; 3638 sweeps averaged)., average of light‐evoked cardiac SNA: note the presence of two peaks (latencies of 63 and 112 ms; control limits: ± 6σ; 1075 sweeps averaged) and the period of inhibition that follows., average of light‐evoked hindlimb MVC SNA evoked by flashing the RVLM region (left); note the presence of two peaks (latencies of 120 and 203 ms; control limits: ± 6σ; 1080 sweeps averaged). On withdrawing the fibre‐optic probe by 2 mm dorsally (right), the faster peak still protrudes above the upper control limit (latency = 122 ms; control limits ± 6σ; 1080 sweeps averaged), whereas the slower peak does not, suggesting that these two peaks arise as a result of stimulation of neurons at different topographical locations. [Color figure can be viewed at wileyonlinelibrary.com

We first studied collateralization of RVLM neurons that innervate anatomically distinct but functionally similar outflows by simultaneously recording MVC activity in the ipsilateral forelimb and hindlimb. Intermittent optogenetic stimulation of RVLM neurons that were retrogradely transduced from the lower spinal cord resulted in time‐locked activation of forelimb SNA (four of four animals), with co‐ activation of hindlimb SNA observed in three of four animals. In all cases, the detected peaks exceeded the background level of nerve activity by >4σ (Fig. 5 A , left) and were abolished by withdrawal of the optrode by 3 mm (Fig. 5 C ). In some cases, smaller stimulus‐locked responses were still apparent when stimulation was delivered 2 mm dorsal to the RVLM (Fig. 6 D ). Such residual activity may reflect the dorsal axonal trajectories of these neurons or their collateralized innervation of the NTS (Lipski et al . 1995 ; Stornetta et al . 2016 ). Peaks in forelimb SNA had response latencies between 125–211 ms, whereas hindlimb SNA peaked at 195–288 ms. Responses were typically single‐peaked, although, in one animal, a double‐peaked response was observed in forelimb MVC activity (peaks at 125 and 211 ms) (Fig. 5 B ). These findings indicate that some RVLM neurons with axonal projections to the lower spinal cord (presumably innervating MVC preganglionic neurons) also provide collateral excitation to upper thoracic preganglionic neurons that supply forelimb MVC.

Examples of stimulus‐triggered averages of sympathetic nerve activity. Dotted lines indicate upper and lower control limits., left: averages of forelimb and hindlimb SNA triggered by light stimulation activation of the RVLM region (473 nm, 10 mW, 20 ms pulse width, 1 Hz; 1501 sweeps averaged); right: averaged nerve responses following withdrawal of the fibreoptic probe by 3 mm (control limits: ± 6σ,= 1468 sweeps)., same average of forelimb SNA depicting a double peak in response to laser activation. After moving the fibreoptic probe 0.3 mm medially, the second peak is no longer present (control limits: ± 4σ, 900 sweeps averaged)., composite drawing showing the location of mCherry‐positive neurons identified in sections that spanned the rostrocaudal extent of the RVLM in the experiment depicted above. [Color figure can be viewed at wileyonlinelibrary.com

, intersectional optogenetic strategy., representative ChR2‐mCherry expression in the RVLM in a successful experiment (scale bar = 100 μm). The relative position of the image is indicated in the camera lucida drawing below., different possible anatomical and functional organizations of collateralized RVLM neurons and the expected result of electrophysiology experiments in each case. SPN, sympathetic preganglionic neuron. [Color figure can be viewed at wileyonlinelibrary.com

Histological sections from 32 rats (nine from experiments in which time‐locked photoactivation of sympathetic nerves was observed; 23 in which it was not) were examined for mCherry fluorescence. mCherry‐labelled RVLM neurons were always observed in experiments in which photostimulation evoked sympathetic responses and tended to be more sparse (14 rats) or absent (9 rats) in experiments during in which no responses were recorded. Labelled neurons from successful experiments were concentrated within the RVLM and, to a lesser extent, the RVMM, with occasional (less than one neuron per experiment) mCherry neurons in the raphé nuclei. No mCherry cells were seen in the C3 or NTS regions (Figs 4 B and 5 C ).

Paired sympathetic nerve recordings were made in 39 rats that had previously received spinal injections of the retrogradely transported vector CAV‐Cre and ipsilateral RVLM injections of a cre‐dependent AAV that drives ChR2. In 13 of these animals, we observed time‐locked activation of one or more sympathetic nerves in response to light pulses delivered to the ipsilateral RVLM.

, ventral view of Waxholm ventral medulla; anatomical landmarks are pyramidal tract (Py), facial nucleus (VII), inferior olive (Inf Ol) and nucleus ambiguus (na)., enlarged view of boxed region indicated in () showing distribution of 273 bulbospinal TH‐immunoreactive neurons, which were used to define the boundaries of the RVLM (green)., positions of spinally projecting neurons; neurons that fell outside of the RVLM boundary defined in () were excluded from the analysis ()., density maps showing 85% boundaries of GFP (green), mCherry (magenta) and double‐labelled (white) RVLM neurons., mean mediolateral (), rostrocaudal () and dorsoventral () co‐ordinates of GFP, mCherry and double‐labelled neurons. Each point shows the mean ± SEM of neurons from one replicate.< 0.01,< 0.001 [Color figure can be viewed at wileyonlinelibrary.com

The spatial distributions of RVLM neurons retrogradely labelled from T 2 , T 10 or both spinal segments were plotted in 3D space and compared (Figs 2 D , E and 3 ). The distributions of all three groups overlapped substantially; however, density maps (Fig. 3 E ) and statistical analysis of the Cartesian co‐ordinates of labelled neurons (Fig. 3 F – H ) indicate that neurons that innervate the upper spinal cord lie caudal and dorsal to neurons that innervate the lower spinal cord. Mean rostrocaudal Waxholm co‐ordinates were 310.5 ± 0.5 voxels (GFP) vs . 314.6 ± 0.7 (mCherry, Tukey's post hoc test, P < 0.001) and 313.8 ± 0.8 (GFP + mCherry, Tukey's post hoc test, P < 0.01) (Fig. 3 G ), corresponding to a mean difference of ∼160 μm. Mean dorsoventral co‐ordinates were 186.5 ± 0.4 (GFP) vs . 184.3 ± 0.6 voxels (mCherry: Tukey's post hoc test, P < 0.01) (Fig. 3 H ), corresponding to a mean difference of 86 μm. There was no significant difference in the dorsoventral co‐ordinates of double‐labelled neurons (185.4 ± 0.5) compared to single‐labelled neurons.

, epifluoresence image of a 50 μm brainstem section at the level of the RVLM showing the distribution of reporters and tyrosine hydroxylase (TH)., experimental scheme: HSV‐GFP and HSV‐mCherry were microinjected at the Tand Tspinal cord, respectively., confocal image of boxed region in (): double‐labelled neurons containing both GFP and mCherry made up ∼20% of the bulbospinal population and included TH‐immunoreactive C1 neurons (, arrow) and TH‐negative non‐C1 neurons (, arrow). Co‐ordinates of bulbospinal neurons were transformed into Waxholm space (shows data corresponding to () aligned onto Waxholm reference dataset., 1561 bulbospinal neurons from 11 replicates projected into 3D Waxholm segmentation model. sp5, spinal trigeminal tract; py, pyramidal tract; na, nucleus ambiguus. [Color figure can be viewed at wileyonlinelibrary.com

We then examined whether colocalization of fluorophores could be observed when HSV variants were microinjected at different spinal segments (HSV‐GFP at T 2 , HSV‐mCherry at T 10 ) (Fig. 2 A – C ). A similar pattern was seen in each experiment; of the 70 ± 8 retrogradely labelled RVLM neurons identified per side (eleven hemi‐medullae from six animals), 21 ± 2% contained both GFP and mCherry, 53 ± 3% contained GFP only (corresponding to labelling from T 2 ) and 26 ± 3% contained mCherry only (corresponding to labelling from T 10 ). Double‐labelled neurons more probably expressed TH than neurons single‐labelled with GFP or mCherry (43% of double‐labelled neurons contained TH compared to 24% of GFP‐ and 29% of mCherry‐only neurons, chi‐squared, P < 0.00001). Similarly, the ratio of double: single‐labelled TH‐positive neurons was higher than for TH‐negative neurons (0.32 vs . 0.18, Fisher's exact test, P < 0.0001). An incidental finding of the current study was the presence of double‐labelled neurons in the rostral ventromedial medulla (RVMM), the midline raphé nuclei, and the C3 cell group, although the prevalence of spinal bifurcation in sympathetic premotor groups other than the RVLM was not systematically investigated.

We first examined whether GFP‐ and mCherry‐driving HSV variants drove equivalent reporter expression by co‐injecting both at the T 2 spinal cord and examining the degree of reporter co‐localization in retrogradely transduced medullary neurons. Labelled neurons were counted in three sections each from two such experiments. The percentage of labelled neurons that contained both reporters was 81% and 91%, respectively (Fig. 1 B ). This suggests that synaptic terminals within the vicinity of the injection site approached saturation, even when vectors were diluted by half, and that retrograde labelling of both was equally efficient.

, reporter expression in the ventral medulla following microinjection of HSV‐hCMV‐GFP at the Tspinal cord. A 930 μm thick optical stack from a CLARITY‐cleared brainstem block; insert shows the corresponding atlas plate at Bregma –12.12 (', enlarged view of region denoted by orange box in ()., control experiment: extensive colocalization of reporter expression following co‐injection of retrograde HSV variants at the Tspinal cord. GFP and mCherry channels are merged such that double‐labelled neurons appear white. Individual channel images shown in () and (). Na, nucleus ambiguus., injection sites in the upper (left) and lower (right) thoracic spinal cord marked by fluorescent beads. [Color figure can be viewed at wileyonlinelibrary.com

Discussion

The present study addresses the organizational principle/s by which RVLM neurons regulate SNA. As discussed in the Introduction, it is clear from the literature that RVLM neurons are not a homogeneous population. The unresolved questions include the extent to which they are anatomically and functionally selective, as well as to what extent the individual RVLM neurons make widespread projections with more generalized sympathetic actions.

Using more refined tools than were previously available, we found that a substantial minority of RVLM neurons send collaterals to diverse spinal segments. Using replication‐deficient retrograde viral tracers injected at upper or lower thoracic spinal segments, we found that, in addition to neurons that expressed only green or red fluorescent reporters (indicating projections to the T 2 or T 10 thoracic cord, respectively), 21% of spinally projecting RVLM neurons were found to contain both.

Next, we used an intersectional approach in which RVLM neurons that project to the lower spinal cord were selectively transduced to express ChR2. Responses to light stimulation of the RVLM were recorded in sympathetic nerves that emerged from both upper and lower spinal cord segments. Intermittent stimulation of retrogradely transduced neurons evoked activity in both upper and lower sympathetic outflows, including both functionally similar (forelimb and hindlimb MVC) and functionally dissimilar sympathetic nerves (cardiac and hindlimb MVC/lumbar sympathetic chain). Electrophysiological responses recorded in rostral outputs occurred at shorter latencies than simultaneously recorded responses recorded in caudal sympathetic outputs. As expected, the latencies of responses evoked in forelimb and cardiac sympathetic nerves were consistent with the latencies of responses evoked by optogenetic or electrical stimulation of RVLM neurons (Morrison & Reis, 1991; Abbott et al. 2009) and with the latencies of antidromic responses to spinal cord stimulation recorded in functionally identified sympathetic premotor neurons (Brown & Guyenet, 1985; Verberne et al. 1999). Although we consider that these results are best explained by activation of RVLM neurons with axon collaterals within the spinal cord, it is worth considering other possibilities.

First, there is the chance that sympathetic outputs from the upper spinal cord were excited not via collateral branches within the spinal cord, but rather by collateral activation of premotor neurons within the RVLM (or other sympathetic premotor nuclei). Anatomical evidence suggests that this is plausible: C1 neurons, including those with spinal axons, give rise to collateral branches that terminate within the ipsi‐ and contralateral RVLM (Lipski et al. 1995; McMullan & Pilowsky, 2012; Turner et al. 2013; Stornetta et al. 2016) and, to a lesser extent, the A5, C3 and raphe pallidus (Stornetta et al. 2016). Furthermore, ultrastructural analysis and trans‐synaptic viral tracing confirms synaptic connectivity between C1 neurons and neighbouring RVLM neurons (both C1 and non‐C1) (Milner et al. 1987; Agassandian et al. 2012; Dempsey et al. 2017). It could therefore be argued that activation of proximal sympathetic outflows in the present study is a result of excitation of untransduced bulbospinal neurons by their ChR2‐expressing neighbours. However, functional studies have yielded no evidence of detectable coupling between bulbospinal neurons in vivo (McAllen et al. 2001) or in vitro (L. Bou Farah & S. McMullan, unpublished observations) and therefore synaptic connectivity between adjacent RVLM neurons is an improbable explanation for our electrophysiological results.

As an alternative explanation, a small portion of preganglionic neurons in the lower thoracic cord that are activated by RVLM stimulation could perhaps project rostrally to innervate sympathetic post‐ganglionic neurons that in turn innervate the forelimb or heart. Similarly, ChR2‐expressing RVLM neurons may target interneurons in the lower cord that innervate preganglionic neurons in the upper thoracic cord. Although it does appear that some preganglionic neurons innervate very distant targets, and can in some cases innervate more than one sympathetic ganglion (Jansen et al. 1993; Pyner & Coote, 1994), the proportion of those that do so is small, even in instances where the ganglia in question are anatomically close and are supplied by neurons with a largely overlapping spinal distribution (Jansen et al. 1993). Moreover, the responses evoked in forelimb or cardiac sympathetic outputs occurred at shorter latencies than those recorded in lumbar or hindlimb outputs in every case in the present study, indicating that responses in proximal outputs were not secondary to a relay in the lower cord.

In the absence of evidence for functional coupling between adjacent medullary bulbospinal neurons or collateralization of sympathetic preganglionic neurons, we conclude that optogenetic activation of premotor sympathetic neurons innervating both upper and lower spinal segments represents the simplest explanation for our observations. Anatomical data from the present study unequivocally demonstrate that such a population of neurons exists. These collateralized axonal projections constitute an entirely plausible anatomical substrate for our electrophysiological observations.

Topographical considerations Our findings are consistent with a study that reported dual‐labelling of RVLM neurons following application of two distinct retrograde trans‐synaptic viruses applied to the stellate ganglion and the adrenal gland of the rat (Jansen et al. 1995). There were sympathetic premotor neurons in the RVLM (and elsewhere) that were infected by both agents. This was considered as evidence indicating that these cells innervate both targets. However, the polysynaptic nature of these viruses, in combination with uncertainty regarding the degree of connectivity between local RVLM sympathetic premotor neurons, makes any interpretation difficult (Morrison, 2001). The results of our anatomical study also share some features with a previous survey in the cat, which used fast blue and Fluoro‐Ruby to retrogradely label RVLM neurons from T 4 and T 10 (Gowen et al. 2012). Although these investigators described a smaller proportion of double‐labelled RVLM bulbospinal neurons (<10%), they found (as did we) that double‐labelling was more common in C1 neurons than in non‐C1 neurons. This finding is consistent with the view espoused by Guyenet et al. (2013) that activation of C1 neurons may underlie the global sympathoexcitation evoked by physiological stressors. However, this does not necessarily translate to a more significant role for C1 neurons compared to non‐C1 neurons in the generation of baseline vasomotor tone, as we have previously argued (Burke et al. 2011). The results of the present study are in stark contrast to those of Tucker and Saper (1985), who found that less than 1% of spinally projecting neurons were double‐labelled when using a combination of fast blue, nuclear yellow or diamidino yellow tracers in the rat. Although differences in the spinal segments chosen for tracer injections could contribute to this discrepancy, it is more probable that differences in the uptake efficiency sensitivities of the tracers used may underlie the higher incidence of double‐labelling observed in the present study. Here, co‐injection of both HSV vectors at the same spinal level led to colocalization of fluorophores in over 80% of neurons. Because Tucker and Saper (1985) found that only 20–30% of spinally projecting neurons were double‐labelled under the same conditions, the true extent of collateralization when those tracers were injected at different spinal levels would have been greatly underestimated. From a technical perspective, the anatomical component of the present study extends the connectomic approach that we recently developed for the analysis, mapping and visualization of anatomical datasets (Dempsey et al. 2017). Specifically, we have now incorporated a density‐mapping technique that plots the anatomical boundaries of a brain structure based on the distribution of the individual neurons that define it. This allowed us to probabilistically define the RVLM based on objective criteria (the distribution of bulbospinal TH‐expressing cells) and to limit our analysis to neurons that fell within this region. The boundary of the RVLM as we have defined it here extends more medially than that described by the widely referred to anatomical atlas of Paxinos and Watson (2006) and more closely resembles a functional map constructed by the examination of cardiovascular responses to medullary glutamate microinjections in the anaesthetized rat (Goodchild & Moon, 2009). Our approach may be useful for other investigators who aim to define the anatomical boundaries of regions of interest using cell‐specific criteria (such as neurochemical phenotype, functional profile, or connectivity). This approach revealed previously unrecognized topographical differences in the distribution of RVLM bulbospinal neurons that project to different spinal cord segments: neurons labelled from the upper spinal cord were located caudal to bifurcating neurons or neurons labelled from the lower spinal cord exclusively.

Limitations A key assumption of our anatomical study is that tracer injected in the upper spinal cord gained entry to neurons forming synaptic terminals in the vicinity of the injection site without labelling fibres of passage. Although labelling of fibres of passage is essentially unavoidable when using traditional chemical tracers (Chen & Aston‐Jones, 1995; Vercelli et al. 2000), this limitation does not appear to apply to HSV vectors, which rely on interactions between the viral envelope glycoprotein complex and cell surface receptors to gain intracellular access (Frampton et al. 2005; Sathiyamoorthy et al. 2017). In neurons, the synaptic adhesion molecule nectin‐1 is the critical mediator of HSV cell entry and infection (Geraghty et al. 1998). Because nectin‐1 is predominantly expressed on presynaptic terminals (Mizoguchi et al. 2002) but is not normally expressed in fibre tracts (Castellanos et al. 2013), we conclude that HSV‐mediated reporter expression probably reflects vector uptake at synaptic terminals. This is probably not the case for the CAV‐Cre used in functional experiments, which appears to be able to retrogradely access neurons via axons of passage as well as synaptic terminals (Schwarz et al. 2015). However, our basic conclusion is unaffected. Vector injections were made only in the lower spinal cord and so, if any passing axons were labelled, they would have been in transit to lumbar or lower levels. Thus, optogenetic activation of upper thoracic sympathetic outflows can confidently be attributed to collateral branches of axons that project to the thoracolumbar cord. It should be noted that the methodology used in the physiology experiments permits the detection of neurons with both upper thoracic and thoracolumbar projections. It does not permit us to definitively identify the functional class of these neurons. We are unable to distinguish between neurons which innervate every studied output (e.g. the vasculature of the forelimb muscle, the heart and the vasculature of the hindlimb muscle) and neurons that innervate only a subset of these targets. Although anatomical and functional studies have demonstrated that the thoracolumbar junction is a locus of preganglionic sympathetic neurons that target muscle vascular beds in the hindlimb (Sonnenschein & Weissman, 1978; Baron et al. 1988), neurons with other targets are also present. Preganglionic sympathetic neurons at this level also innervate the inferior and, to a lesser extent, the superior mesenteric ganglion (Strack et al. 1988) and also target the pelvic viscera and associated vasculature (Sonnenschein & Weissman, 1978; Kaleczyc, 1998; Hsieh et al. 2000). Thus, although our anatomical data indicate that collateralization of sympathetic premotor neurons is widespread, our electrophysiological observations do not extend further than the pairs of outflows that we studied. Finally, the stimulus‐triggered averaging used to detect laser‐triggered responses in sympathetic outputs warrants consideration. This commonly used technique has the benefit of high sensitivity, enabling the pooling of thousands of responses to repeated stimuli to distinguish small excitatory or inhibitory events from background noise. However, the amplitude of these responses is greatly influenced not only by the number of active fibres within the nerve, but also by factors that are impossible to control and probably vary from experiment to experiment: these include the number of inactive or damaged fibres, the amount of fat or connective tissue in the nerve sheath, the hydration of the nerve and the properties of the recording electrodes. Therefore, this method cannot be readily used to compare the amplitudes of responses recorded in different animals: it should be viewed as a sensitive tool for the detection of collateralization but with limited use for its quantification. The comparison of responses that were simultaneously recorded from rostral and caudal sympathetic outputs would be similarly limited. Responses recorded from rostral outputs (e.g. forelimb muscle vasoconstrictor) in response to optogenetic stimulation are presumably the result of activation of RVLM neurons that project to the caudal spinal cord but that possess axon collaterals that innervate the rostral spinal cord. However, because responses recorded from caudal outputs (e.g. hindlimb muscle vasoconstrictor) are driven by RVLM neurons with terminals at the same spinal level to which the retrograde viral vector was applied (the caudal spinal cord), the responses recorded from these outputs are presumably driven by collateralized and non‐collateralized neurons alike.