Chemogenetic activation of the diaphragm after spinal cord injury in rats Journal: Respiratory Physiology and Neurobiology

Abstract

We tested the hypothesis that activation of DREADDs in the mid-cervical spinal cord could restore diaphragm activation during spontaneous breathing after cervical spinal cord injury (SCI). Adult Sprague Dawley rats (n = 7) received bilateral mid-cervical ventral horn injections of an AAV construct encoding an excitatory DREADD (AAV9-hSyn-HA-hM3D(Gq)-mCherry; titer: 2.44×10¹³ vg/mL). Subsequently, diaphragm electromyogram (EMG) activity was recorded during spontaneous breathing under isoflurane anesthesia. The selective DREADD ligand JHU37160 (J60) was administered intravenously at acute (3 days), sub-acute (2 weeks), and chronic (2 months) timepoints following cervical hemilesion at spinal level C2. J60 administration resulted in robust increases in diaphragm EMG output at all timepoints, and near-complete restoration of diaphragm EMG activity from the paralyzed hemi-diaphragm in 50% of trials. Administration of J60 to DREADD naïve, spinal intact rats (n = 8) did not produce an increase in diaphragm activity. These proof-of-concept results indicate that refinement of this technique may provide a strategy for improving diaphragm activation after cervical SCI.

Introduction

The diaphragm serves as the primary inspiratory muscle and is innervated by phrenic motoneurons in the C3-C5 spinal cord¹. Cervical spinal cord injury (SCI) typically results in diaphragm paresis or paralysis which directly contributes to impaired breathing. In severe cases, this necessitates mechanical ventilation or diaphragm pacing² as cervical SCI almost always leads to reduced lung volumes and weakened ability to cough³.

Designer receptors exclusively activated by designer drugs (DREADDs) provide a tool to selectively activate or inhibit neurons⁴. In regards to breathing, DREADDs can effectively increase respiratory-related motoneuron discharge as shown by studies of the hypoglossal motor system⁵. For example, when an excitatory DREADD is expressed in hypoglossal motoneurons, subsequent ligand delivery can increase tongue motor output^6–9 and dilate the pharynx⁸. We recently used a ChAT-Cre rat model to target excitatory DREADD expression to phrenic motoneurons. The selective DREADD ligand JHU37160¹⁰ (J60) was able to produce a large increase in diaphragm activation which corresponded to increased inspiratory tidal volume during spontaneous breathing^11,12. Other studies have shown that activation of interneurons in the mid-cervical spinal cord can produce an increase in diaphragm activity in spontaneously breathing mice¹³. Therefore, mid-cervical motoneuron- and interneuron-targeted DREADD expression can be used to stimulate diaphragm activation and breathing.

Here we determined if expression of an excitatory DREADD in the mid-cervical spinal cord could stimulate diaphragm activity during spontaneous breathing in adult Sprague-Dawley rats following cervical SCI. An adeno-associated viral vector (AAV) construct encoding the hM3D(Gq) DREADD was delivered via intraspinal microinjection, targeting the C4 ventral horns. We sought to determine if an AAV approach that generally targeted phrenic motoneurons, but would also likely produce expression in other neurons in the region, would enable DREADD-mediated activation of the diaphragm after SCI.

Rats were implanted with indwelling diaphragm electrodes so that electromyogram (EMG) activity could be recorded and compared across days to weeks post-SCI. The repeated measures design allowed us to determine if the response to the DREADD ligand would wane over time post-injury. These proof-of-concept experiments demonstrate inducing expression of an excitatory DREADD in the mid-cervical spinal cord enables the DREADD ligand, J60, to increase diaphragm activation, with efficacy lasting up to several months after cervical SCI.

Methods

Experiments were carried out using adult Sprague Dawley rats (Envigo; n = 15; females = 8; 2 months old ± 2 weeks at time of first procedure; IACUC protocol #202107438). Animals were housed individually in a controlled environment with a 12-hour light-dark cycle and access to food and water ad libitum. All experiments were conducted in accordance with the NIH Guidelines Concerning the Care and Use of Laboratory Animals and were approved by the University of Florida Institutional Animal Care and Use Committee.

Adeno-Associated Viral Vectors

Rats (n = 7; females = 4) underwent intraspinal injections (see section below) of an AAV vector encoding the excitatory DREADD (hM3D(Gq)) under a human synapsin promoter (AAV9-hSyn-HA-hM3D(Gq)-mCherry; titer: 2.44×10¹³ vg/mL). The pAAV-hSyn-hM3D(Gq)-mCherry transgene plasmid used to produce the vector product was gifted from the laboratory of Dr. Brian Roth at the University of North Carolina (Addgene plasmid # 50474; http://n2t.net/addgene:50474; RRID: Addgene_50474). The viral preparation was generated and titered by the University of Florida Powell Gene Therapy Center Vector Core Lab. Vectors were purified by iodixanol gradient centrifugation and anion-exchange chromatography as previously described¹⁴. Viral aliquots were stored at −20°C and thawed immediately before first use. Any remaining virus was stored at 4°C.

Intraspinal Injections

For these proof-of-concept experiments, the AAV construct was delivered prior to the SCI, such that spinal DREADD expression would be evident before injury. Rats were anesthetized using ketamine (100 mg/kg) and xylazine (10 mg/kg) delivered intraperitoneally. Animals were placed prone on a circulating water heating pad, and a midline incision was made at the base of the skull extending caudally to the shoulder blades revealing the underlying musculature. The muscles of the back were cut and separated at midline from approximately spinal segment C1 to C7. A micro-curette was used to clear the paravertebral muscles and connective tissue overlaying lamina C3-C5. Using a pair of rongeurs, the entire dorsal C4 lamina was removed exposing the dura mater. After bilateral durotomy at C4, rats were moved to a stereotaxic frame (Kopf) for intraspinal injections. A Hamilton syringe (34-gauge needle) was used to inject 1 µl of AAV bilaterally into the ventral horns at C4 (Figure 1; injection coordinates: 0.7 mm lateral to midline at a depth of 1 mm)¹⁵. Following each injection, the needle was left in place for 5 minutes. The overlying muscle and fascia were then sutured with absorbable suture, and the skin closed. Animals were maintained in temperature-controlled incubators until they recovered from anesthesia and then were returned to their home cage. Animals received a post-operative analgesia regiment of subcutaneous buprenorphine (1 mg/kg; extended-release formulation; given once post-operatively) and carprofen (5 mg/kg, q.d.) for the first 48 hours post-surgery.

Figure 1. Intraspinal injections of AAV9-hSyn-HA-hM3D(Gq)-mCherry drives transgene expression in mid-cervical neurons.

Schematic of bilateral intraspinal injections (left) of AAV9-hSyn-HA-hM3D(Gq)-mCherry into the C4 ventral horns targeting the phrenic motor nuclei (red columns). The call-out photomicrograph shows example histology from the C4 ventral horn. Neurons transduced with the hM3D(Gq) DREADD appear red.

Diaphragm EMG Lead Implantation

Four weeks following intraspinal AAV injections, animals underwent an additional procedure to implant pairs of indwelling EMG electrodes bilaterally into the mid-costal diaphragm¹⁶. Animals were anesthetized and maintained during the surgical procedure as described in the previous section (see Intraspinal Injections). After an adequate depth of anesthesia was reached, animals were placed prone in a Kopf stereotaxic frame over a heating pad. A midline incision was made in the skin covering the dorsal aspect of the skull revealing the skull surface from the frontal bone to the lambda suture. Six screws were placed in the skull to serve as anchors for the head cap. Two screws were placed in the left and right parietal bones, respectively, with two more placed in the frontal bone. A ground wire was wrapped around the three screws on the left portion of the skull. The head cap was then secured to the anchoring screws using dental acrylic. After the head cap was fully set, the animal was removed from the stereotaxic frame. Subsequently, EMG leads attached to the head cap were subcutaneously tunneled down the animal's back and externalized through a small incision in the skin overlying their abdominal muscles. A midline laparotomy was performed to access the diaphragm. Pairs of electrode wires (stainless steel; AS631, Cooner Wire Inc., Chatsworth, CA) were placed in each hemi-diaphragm. Wires in each electrode pair were placed approximately 3 mm apart and were secured into the diaphragm with a surgical knot. Following EMG lead implantation, the abdominal muscle and overlying skin were sutured closed. Rats were maintained in temperature-controlled incubators until they recovered from anesthesia. The postoperative analgesia regimen included subcutaneous buprenorphine (1 mg/kg; extended-release formulation; given once postoperatively) and carprofen (5 mg/kg, q.d.) for the first 48 hours post-surgery.

SCI Surgery

Hemisection of the C2 spinal cord (C2Hx) was performed as described previously^16,17. Rats were anesthetized with ketamine/xylazine and placed supine on a circulating water heating pad to maintain body temperature. Under aseptic conditions, a dorsal, midline incision was made from the base of the skull to approximately spinal level C4. The underlying neck musculature was cut and separated at midline revealing the spinal laminae. Using a micro-curette the dorsal paravertebral muscles between C1 and C3 were cut away and retracted revealing the laminae bones. The left caudal portion of the C2 lamina was removed, leaving the underlying dura mater intact. A micro-knife (Cat # 10055–12, Fine Science Tools) was used to perform a hemisection injury on the left side of the spinal cord. The micro-knife was inserted at midline and moved laterally. This process was repeated twice to ensure complete injury. The overlying muscles and skin were sutured, and rats were then placed in temperature-controlled incubators. The postoperative analgesia regimen included subcutaneous buprenorphine (1 mg/kg; extended-release formulation; given once post-operatively), carprofen (5 mg/kg, q.d.), and lactated ringers (6 mL/day, q.d.) for the first three days after surgery.

Diaphragm EMG Recordings

Diaphragm EMG recordings were performed 1-day before C2Hx, and then at 3 days, 17 days, and 2 months post-injury. The EMG recording protocol was as follows. Anesthesia was induced with 3% isoflurane in 100% O₂. Once the hindlimb withdrawal reflex was abolished, rats were moved onto a closed-loop electric heating pad to maintain body temperature at 37 ± 0.5°C. A tail vein catheter was placed to allow for administration of the DREADD ligand and vehicle. Diaphragm EMG signals were recorded via a cable plugged into the head cap. Chronic implantation of EMG leads enabled multiple recordings from each rat (i.e., repeated measures design). EMG signals were amplified (1000x) and filtered (100–1000 Hz) using a differential amplifier (A–M systems model 1700). Signals were digitized at 10 kS/s using a Power 1401 (CED, Cambridge, UK). Once a stable plane of anesthesia was reached, rats underwent a 10-minute recording to establish baseline diaphragm EMG parameters. Subsequently, rats received infusions of vehicle (0.6 ml of saline infused over 2 minutes, IV) followed by a 30-minute recording. Rats then received an IV infusion of the selective DREADD agonist, J60¹⁰ (0.1 mg/kg diluted to 0.6 ml final volume, HB6261, HelloBio), and recordings continued for an additional 30 minutes. At the conclusion of each experiment, rats were returned to their home cage after recovering from anesthesia.

J60 Control Experiments

Spinal intact adult Sprague Dawley rats (n = 8; n = 4 females) were studied to assess the impact of the J60 ligand (0.1 mg/kg) on diaphragm EMG activity when no DREADD expression was present (i.e., “DREADD naïve” rats). Anesthesia was induced with 3% isoflurane in 100% O₂ and rats were then placed on a heating pad that maintained body temperature at 37 ± 0.5 °C. Anesthesia was maintained (2–2.5% isoflurane) via a nose cone. An IV catheter was placed to allow administration of vehicle (saline, 0.6 mL) and J60 (0.1 mg/kg, 0.6 mL). Rats were placed supine and after laparotomy de-insulted tungsten wires (127 µm diameter; A-M Systems) were placed into the mid-costal diaphragm bilaterally. Following confirmation of EMG signal, rats were placed supine for the remainder of the experiment. The protocol and recording parameters (e.g., filters, sampling rate) used in these studies of DREADD naïve rats were identical to those used in rats with spinal cord DREADD expression (see Diaphragm EMG Recordings). At the conclusion of the experiment, rats were euthanized with pentobarbital sodium and phenytoin sodium (150 mg/kg, IV). Death was confirmed via thoracotomy.

Histology

Following the final recording timepoints rats in the experimental group (i.e., spinal DREADD expressing animals) were euthanized via transcardial perfusion with heparinized saline followed by 4% paraformaldehyde. Spinal cords were then harvested and post-fixed for 24 hours in 4% paraformaldehyde and then moved to a cryo-protecting solution (30% sucrose in 1x PBS) for a minimum of three days. Cervical spinal cords were blocked in optimal cutting temperature media, cryosectioned longitudinally at 20 µm, and cover slipped. A mCherry fluorescent tag fused to the hM3D(Gq) DREADD allowed for visualization of transduced cells via fluorescence microscopy.

Data Analysis and Statistics

Diaphragm EMG signals were analyzed using a custom MATLAB script (The Mathworks, Natick, MA) to determine average EMG burst peak amplitude, tonic activity, and respiratory rate. The EMG waveforms were digitally filtered using a 2^nd order, 100–1000 Hz bandpass Butterworth filter. The filtered waveforms were rectified by taking the absolute value of the signal, then integrated using a moving median filter with a 150 ms time constant, followed by a moving average filter also with a 150 ms time constant. Phasic (inspiratory) activity in the diaphragm could be detected as the peak EMG burst amplitude. Peak amplitude was determined using the built-in MATLAB “findpeaks” function. Discharge of diaphragm motor units was also present across the expiratory phase of the respiratory cycle, and we considered this to be “tonic” diaphragm activation. To quantify this tonic activity, we determined the minimum amplitude of the EMG trace between phasic EMG bursts. Specifically, we segmented the EMG bursts and surrounding expiratory regions, capturing a window that spanned 550 ms preceding the peak sample to the 550 ms following the peak sample. The minimum amplitude within each segment was then determined using MATLAB’s built-in “min” function. The average respiratory rate was determined by taking the number of detected peaks in a given data section and dividing it by the length of the trace in minutes.

Normalization procedures can be problematic when describing EMG bursting after SCI¹⁸. For example, “% baseline” is not possible when baseline = 0, and low baseline output (e.g., a few active motor units) often results in exceedingly large values when normalized (e.g., thousand-fold increase over baseline). For these reasons, we elected to present the data in absolute values (i.e., mV) as recorded during the experimental procedure. We feel that this approach provides the most straightforward presentation of the data and the response to the DREADD ligand.

The JMP statistical software (version 14.1.0, SAS Institute Inc., Cary, NC) was used to run restricted maximum likelihood (REML) repeated measures mixed linear model analyses on all variables. Only three rats were studied at the two-month timepoints, and these data were therefore not included in the statistical analyses. The results are included as supplemental data, however, as we feel they do provide additional insight into the sustained ability of DREADDs to activate the diaphragm. For significant main effects with more than two levels, the Tukey HSD post hoc test was used to further investigate any significant main effects. Student’s t was used for significant main effects with only two levels. For all statistical analyses α = 0.05.

Results

Impact of DREADD activation on diaphragm EMG activity

Application of the J60 ligand resulted in robust, bilateral increases in diaphragm EMG activity. Figure 2 provides a representative example of J60-induced EMG activation at 3 days post-SCI.

Figure 2. Impact of DREADD activation on diaphragm EMG 3 days post-cervical SCI.

Example traces of left (a) and right (b) rectified and integrated diaphragm EMG activity from a spinal injured Sprague Dawley rat that had previously undergone injections of AAV9-hSyn-HA-hM3D(Gq)-mCherry into the C4 ventral horns before and after application of the DREADD ligand, J60. Callout panels below each trace show example activity at baseline (a_i, b_i), after injection of vehicle (a_ii, b_ii), and after application of the DREADD ligand, J60 (a_iii, b_iii).

Activity in the left diaphragm (i.e., ipsilateral to the C2Hx) was attenuated or absent after the injury. This can be appreciated in Figure 3a, in which the vehicle (saline) treated condition has little to no burst amplitude in the left diaphragm at 3- and 17-days post-injury. The J60 ligand produced a considerable increase in left diaphragm EMG bursting at all timepoints (treatment effect, p < 0.001; Supplemental Table 1).

Figure 3. DREADD activation increases diaphragm EMG peak amplitude and decreases respiratory rate in rats following cervical SCI.

Plots depict the average peak amplitude of rectified and integrated diaphragm EMG bursts for the left (a) and right (b) hemi-diaphragm along with respiratory rate (c) following administration of vehicle (blue) and J60 (orange) at pre-injury, 3 days post-injury, and 17 days post-injury. Scatter points depict average values for each animal. Statistical reports for all panels are provided in Supplemental Table 1. Bpm = breaths per minute, dia = diaphragm, EMG = electromyography, mV = milli-volts, pk amp = peak amplitude.

We did not detect post-injury alterations in the EMG burst amplitude in the right hemi-diaphragm (i.e., contralateral to C2Hx) Figure 3b, Supplemental Table 1). However, there was a significant increase in the right diaphragm EMG burst amplitude following application of the J60 ligand (treatment effect, p = 0.002; Supplemental Table 1).

The impact of DREADD activation on the inspiratory burst rate (per minute) of the EMG signal is shown in Figure 3c. Delivery of the J60 ligand resulted in a decrease in respiratory rate as compared to the vehicle treatment (treatment effect, p < 0.001; Supplemental Table 1). Thus, the increased inspiratory EMG burst amplitude (Fig. 3a-b) was associated with a decrease in the corresponding inspiratory burst rate.

Application of the J60 ligand also resulted in an increase in tonic diaphragm EMG activity (Figure 4). This effect was statistically significant in the left diaphragm (treatment effect, p = 0.004; Supplemental Table 1), with a strong suggestion in the data that the response was greater after SCI (vs. spinal intact). The apparent increase in the tonic response in the right diaphragm after the DREADD ligand was delivered did not reach statistical significance (treatment effect, p = 0.079; Supplemental Table 1).

Figure 4. DREADD activation increases diaphragm tonic activity in rats following cervical SCI.

Plots depict the average tonic activity of rectified and integrated diaphragm EMG bursts for the left (a) and right (b) hemi-diaphragm following administration of vehicle (blue) and J60 (orange) at pre-injury, 3 days post-injury, and 17 days post-injury. Scatter points depict average values for each animal. Statistical reports for all panels are provided in Supplemental Table 1. Dia = diaphragm, mV = milli-volts.

In a few additional experiments, we recorded the left diaphragm EMG response to J60 at two months following the C2Hx injury (Supplemental Figure 1). The remaining sample (n = 3) was too low to be included in the overall mixed linear model analyses that are presented in Figures 3–4. However, the data are informative and suggest that the ability of the DREADD ligand to produce increased diaphragm output is maintained in the chronic SCI time frame. As shown in Supplemental Figure 1, a strong increase in EMG output in response to J60 is present in 2 of 3 rats at 2 months following C2Hx.

DREADD Ligand Control Experiments

The impact of the DREADD ligand (J60) on diaphragm EMG activity was evaluated in spinal intact, DREADD naïve rats (n = 8) (Supplemental Figure 2). EMG peak amplitude was not impacted by the sham (saline) IV injection (Supplemental Figure 2a). In sharp contrast to the response observed in rats with DREADD expression (Figures 2–3), administration of the J60 ligand produced an ~20% reduction in diaphragm EMG burst amplitude (left hemi-diaphragm: − 20.3±16.4%; right hemi-diaphragm: −22.9±18.5 %). However, this apparent decrease was not statistically significance as evaluated using REML mixed linear model (timepoint effect, p = 0.1563; Supplemental Table 2). Diaphragm tonic activity (Supplemental Figure 2b) was consistent across all timepoints, with no change following J60 application (timepoint effect, p = 0.9949; Supplemental Table 2). The respiratory rate (Supplemental Figure 2c) declined after application of J60 (timepoint effect, p < 0.0001; Supplemental Table 2).

Histology

Post-mortem histological analysis of the cervical spinal cord was completed in all rats in the experiment group. In addition to the DREADD, the AAV construct also drove expression of a mCherry fluorescent protein, which served as a marker of AAV transduction. Histology of control rats was not assessed as these animals were vector naïve and therefore would not show evidence of fluorophore expression. Expression of mCherry was detected by fluorescent microscopy in the mid-cervical spinal cord of all rats (i.e., 7 of 7 experimental rats); an example is provided in Figure 1. The overall mCherry expression pattern was diffuse, extending from spinal segment C3 to C7 in all rats. Qualitative evaluation with microscopy confirmed widespread mCherry expression in neurons throughout the mid-cervical spinal cord, with the greatest density in the ventral horn. Further, mCherry expression was present on both sides of the spinal cord in all rats. The histological processing methods did not allow correlation of the relative degree of mCherry expression with the diaphragm EMG data.

Discussion

We demonstrated that expressing an excitatory DREADD in the mid-cervical spinal cord enables the high affinity DREADD ligand J60 to robustly activate the diaphragm after cervical SCI. The response to J60 was sustained for up to 30 minutes and could be effectively induced at acute through chronic post-injury timepoints. The data build upon prior reports targeting DREADDs to propriospinal neurons^13,19, further supporting the feasibility of using chemogenetic methods to increase diaphragm motor output following cervical SCI.

Breathing after Cervical Spinal Cord Injury

Respiratory impairments are common following cervical SCI^2,3,20–22. The alpha-motoneurons that innervate the diaphragm, and intercostal muscles are located in the cervical and thoracic spinal cord, respectively. Damage to the cervical cord can result in the destruction of these neurons or the axon tracts connecting respiratory motor nuclei with the brainstem. The extent of respiratory impairment after SCI is dependent on the level and severity of the injury, with C3-C4 lesions typically leading to the most severe impairments^21,22. High cervical SCI patients often require full-time respiratory support via mechanical ventilation. With more caudal lesions, individuals may be able to breathe independently but struggle with higher-force respiratory behaviors such as coughing and clearing their airways². Complications such as pneumonia, atelectasis, and respiratory insufficiency are common due to reduced tidal volumes and the inability to effectively clear their airways^20,22. The clinical standard of care consists of respiratory support and therapy to preserve alveolar ventilation and prevent infection². However, these therapeutics are not sufficient for improving respiratory function in all cases. Therefore, there is a need to develop new methods to improve respiratory muscle function following cervical SCI.

Neuromodulatory Methods to Improve Breathing after Cervical SCI

Recently, several neuromodulatory methods have been developed that show promise for improving respiratory muscle function after cervical SCI. Phrenic long-term facilitation (pLTF) following acute intermittent hypoxia (AIH) exposure is one of the most widely studied forms of respiratory neural plasticity^23–25. In short, exposure to brief, repeated bouts of hypoxia results in a sustained increase in phrenic motor output that persists for several hours following exposure²⁶. One mechanism of pLTF evoked by relatively modest hypoxia levels involves the release of serotonin onto phrenic motoneurons from serotonergic neurons in the raphe nucleus²⁶. Serotonin initiates new BDNF synthesis via protein kinase C, which activates tropomyosin receptor kinase B receptors (TrkB) on phrenic motoneurons, subsequently activating ERK MAP kinases.²⁷ It is speculated that ERK MAP kinases play a role in phosphorylating phrenic glutamate receptors, which strengthens the neural connections between pre-motor respiratory areas and phrenic motoneurons, ultimately enhancing phrenic motor output²⁸. Pre-clinical studies have firmly established that AIH treatments can be used to improve diaphragm output in rodent models of cervical SCI^29–32. AIH will also induce a long-term facilitation of ventilation in humans with SCI³³. This discovery has prompted clinical trials assessing AIH as a therapeutic to improve respiratory and non-respiratory motor function in individuals living with SCI. These trials have demonstrated that AIH is safe and well tolerated by individuals with spinal cord injury³⁴ and can lead to improvements in respiratory^33,35 and other somatic motor function^36–39.

Positive allosteric modulators of AMPA receptors, also known as ampakines, are another neuromodulatory method capable of improving diaphragm muscle activation following SCI. As allosteric modulators, ampakines bind to secondary binding sites on AMPA receptors, inducing conformational changes that enhance AMPA currents when the primary ligand, glutamate binds to the receptor⁴⁰. AMPA receptors are robustly expressed on phrenic motoneurons ^41,42 leading to the investigation of ampakines as a potential modulatory method for increasing phrenic output. Proof-of-concept work has demonstrated that the application of ampakines in spinal intact rodents results in sustained increases in phrenic motor output^43,44. Further work in a rodent model of cervical SCI has demonstrated that ampakines are effective for improving phrenic output in anesthetized rodents⁴⁵ and can improve diaphragm function and tidal volume in awake freely behaving rats post-injury¹⁶. Currently, there are no ongoing clinical trials using ampakines to treat individuals with SCI. However, a number of clinical studies have demonstrated that ampakines are generally safe and feasible for use in humans^46–50. Therefore, ampakine treatment may offer a promising avenue of treatment for individuals with respiratory muscle dysfunction following SCI.

Other relevant techniques include the use of viral vectors to induce expression of optogenetic receptors⁵¹ and the neural growth factor brain-derived neurotrophic factor (BDNF)⁵² in the area of the phrenic nuclei. Alilain et al. demonstrated that viral-mediated expression of the light-activated cation channel, channelrhodopsin-2 (ChR2) in the area of the phrenic nuclei was effective for restoring diaphragm output during photostimulation following C2Hx injury⁵¹. This technique also appeared to evoke a form of spinal plasticity in the phrenic neural circuitry as increased diaphragm output persisted following cessation of photostimulation⁵¹. Work from others has demonstrated that AAV mediated overexpression of BDNF and/or its neuronal receptor, TrkB at the level of the phrenic motor pool can restore diaphragm activity after a cervical SCI^52–54.

DREADD Activation of Spinal Neurons

DREADDs have been used extensively to study neural circuits across the brain and spinal cord^4,55. Pharmacologically selective actuator module (PSAM), a technology similar to DREADDs, can effectively activate lumbar motoneurons in mouse models of amyotrophic lateral sclerosis (ALS)^56,57. Other work has shown that expression of excitatory DREADDs in spinal motoneurons can help facilitate peripheral nerve regeneration following sciatic nerve injury^58,59. Previous work from our group demonstrated that activation of excitatory DREADDs in mid-cervical spinal cord neurons can effectively increase diaphragm output¹². Using a ChAT-Cre (spinal intact) rat model, we found that targeting DREADD expression to mid-cervical motoneurons – targeting the phrenic motor pool - enabled the J60 ligand to increase phrenic motor output and inspiratory tidal volume. We speculate that this spinally directed chemogenetic approach could be developed into a viable strategy for enhancing tidal volume in cases of partial diaphragm paralysis following incomplete cervical SCI, potentially allowing alveolar ventilation to be maintained without additional ventilatory assistance.

In regard to interneuron populations, Jensen et al. showed that DREADD-mediated activation of spinal V2a interneurons restored diaphragm activity following cervical SCI¹³. They found that selective activation of V2a interneurons using the excitatory hM3D(Gq) DREADD was able to restore diaphragm activity at acute through subacute timepoints after a C2Hx lesion¹³. Further, inhibition of V2a neurons impaired diaphragm activation when mice were exposed to a maximal chemosensory gas challenge¹³. The V2a spinal interneurons are glutamatergic and reside in the middle spinal laminae. Anatomical tracing confirms direct synaptic connections between V2a interneurons and phrenic motoneurons, and such connections may increase after SCI⁶⁰. Collectively, these prior studies establish a role for V2a interneurons in the recovery of diaphragm activity following cervical SCI in mouse models and demonstrate the potential of DREADD-based approaches to enhance diaphragm activation post-injury. Building upon this foundation, we show that using AAV to induce non-specific DREADD expression in the rat spinal cord can drive sustained increases in diaphragm output at acute through chronic timepoints after cervical SCI.

Neural Substrate Underlying the Diaphragm Response to Spinal DREADD Activation

The most straightforward explanation of the observed increase in diaphragm EMG bursting following J60 administration is direct phrenic motoneuron activation¹². Several prior studies that have selectively expressed DREADDs in hypoglossal motoneurons confirm the effectiveness of the method for activating respiratory-related discharge^6–9. Alternatively, DREADD-mediated stimulation of excitatory interneuron populations that synapse on phrenic motoneurons may have occurred¹³. Propriospinal neurons in the mid-cervical spinal cord provide direct synaptic inputs to phrenic motoneurons⁶¹, and activation of spinal interneurons can directly modulate phrenic output⁶². Further, activation of propriospinal neurons can increase diaphragm activity following cervical SCI in rodents^13,61,63,64.

In this current work, we noted widespread neuronal expression of the mCherry marker (and by extension, hM3D(Gq)) throughout the cervical spinal cord. It is not possible to directly confirm the underlying neuronal substrate, but the DREADD-mediated increase in diaphragm output was likely driven by a combination of phrenic motoneuron and propriospinal neuron activation. However, the non-specific method of AAV delivery used here (i.e., likely transducing both propriospinal neurons and phrenic motoneurons) is not sufficient to limit transduction to the phrenic system and therefore, is prone to off-target effects. In preliminary studies using spinal injections of AAV9-hSyn-HA-hM3D(Gq)-mCherry, we observed sporadic forelimb movements when non-anesthetized rats were given J60. This was likely due to unintentional transduction of mid-cervical spinal moto- and interneuron circuits that innervate the muscles of the forelimbs. Since the current study was designed as a proof-of-concept trial, we elected to study the rats under isoflurane anesthesia to enable high-quality diaphragm EMG recordings under more controlled conditions. Under these conditions, we did not observe forelimb movements or other obvious off-target responses. In previous experiments, we were able to achieve DREADD-mediated diaphragm activation in awake freely behaving animals without any detectable off-target forelimb movements¹². However, this required the use of ChAT-Cre transgenic animals along with Cre-dependent viral constructs that ultimately allowed us to limit DREADD expression primarily to the phrenic motor pools. Therefore, for this technique to be feasible in wildtype animals AAV delivery techniques need to be refined to limit expression of DREADDs to phrenic motoneurons.

Selectivity of the J60 ligand

JHU37160 (i.e., J60) is a DREADD ligand that readily crosses the blood brain barrier, making it advantageous for CNS applications compared to tradition DREADD ligands such as clozapine-N-oxide, which are generally considered to have poor brain penetrance⁶⁵. As a relatively new ligand, the off-target profile of J60 is actively being evaluated. The original work describing J60 reported an effective range for DREADD activation with minimal off-target effects at 0.01–1 mg/kg for mice and 0.01–0.3 mg/kg in rats¹⁰. Subsequent studies have reported no observable off-target effects at 0.1⁶⁶, 0.03⁶⁷ and 1 mg/kg⁶⁸ in C57/bl6 HSD mice, wildtype C57/bl6 mice, and vGluT2-Cre mice, respectively. Similarly, no off-target impact was described following a 0.2 mg/kg J60 dose in Long Evans rats⁶⁹. Only a few prior studies have assessed the impact on J60 in the context of breathing and respiratory muscles. One study assessing a chemogenetic method to increase genioglossus muscle activation and upper airway patency observed no detectable effect of the J60 ligand on tongue EMG activity in sham vector treated mice⁹. An additional study found no effect of J60 (0.1 mg/kg; IP) on ventilation or maximum inspiratory flow in DREADD naïve Lepr^b -Cre mice ⁷⁰.

Our control experiments, in contrast to the aforementioned studies, indicate that J60 may have a biologic impact on the neural control of breathing, independent of DREADD expression. Indeed, we observed a statistically significant decrease in respiratory rate following intravenous J60 administration (0.1 mg/kg) in spinal-intact, vector-naïve rats. This was accompanied by a trend toward reduced diaphragm EMG amplitude. We conclude that the 0.1 mg/kg J60 dose is having an impact on respiratory output. Ultimately, we do not believe this confounds our main finding that mid-cervical DREADD activation robustly increases diaphragm output following SCI. The robust increase in diaphragm EMG output that was induced by J60 in spinally injured rats with spinal DREADD expression shows that the direct spinal effect of J60 in this condition was much more powerful than any centrally mediated inhibition. Nevertheless, we suggest that future DREADD studies evaluating respiratory outcomes should consider using lower J60 doses (i.e., < 0.1 mg/kg) which may mitigate the off-target effects on respiration. Additionally, alternative DREADD ligands could be explored to identify ligands with minimal respiratory impact.

Conclusions

These proof-of-concept experiments demonstrate that activating excitatory DREADDs expressed in neurons of the mid-cervical spinal cord is effective for improving diaphragm activation following cervical SCI. DREADD-mediated enhancement of diaphragm output was observed at multiple acute to chronic timepoints post-injury, demonstrating this technique's long-term feasibility. Even without specific targeting of the phrenic motoneuron pool, the J60 ligand triggered robust increases in diaphragm activation after SCI. Moving forward, however, we suggest that directly targeting DREADD expression to phrenic motoneurons will be necessary to minimize off-target activation. This may be possible via retrograde viral transport following vector delivery to the phrenic nerve or diaphragm muscle since this approach is highly effective in the hypoglossal motor system^71,72. Intersectional genetics methods may also enable selective targeting of phrenic motoneurons as recently described⁷³.

Highlights.

• Respiratory problems are common after cervical spinal cord injury.

• The diaphragm serves as the primary inspiratory muscle and is suspectable to impaired activation following cervical spinal cord injury.

• Here we developed a novel chemogenetic method to improve diaphragm activation following cervical spinal cord injury.

Abbreviations:

AAV

adeno-associated virus

AIH

acute intermittent hypoxia

BDNF

brain-derived neurotrophic factor

C2Hx

C2 hemisection

DREADD

designer receptors exclusively activated by designer drugs

EMG

electromyography

IV

intravenous

J60

JHU37160

pLFT

phrenic long term facilitation

REML

restricted maximum likelihood

SCI

spinal cord injury

TrkB

tropomyosin receptor kinase B

Data and Code Availability:

The datasets along with MATLAB code generated during the current study are available from the corresponding author upon reasonable request.