Phrenic motor neuron loss in an animal model of early onset hypertonia

Abstract

Phrenic motor neuron (PhMN) development in early onset hypertonia is poorly understood. Respiratory disorders are one of the leading causes of morbidity and mortality in individuals with early onset hypertonia, such as cerebral palsy (CP), but they are largely overshadowed by a focus on physical function in this condition. Furthermore, while the brain is the focus of CP research, motor neurons, via the motor unit and neurotransmitter signaling, are the targets in clinical interventions for hypertonia. Furthermore, critical periods of spinal cord and motor unit development also coincide with the timing that the supposed brain injury occurs in CP. Using an animal model of early-onset spasticity (spa mouse [B6.Cg-Glrbspa/J] with a glycine receptor mutation), we hypothesized that removal of effective glycinergic neurotransmitter inputs to PhMNs during development will result in fewer PhMNs and reduced PhMN somal size at maturity. Adult spa (Glrb−/−), and wild-type (Glrb+/+) mice underwent unilateral retrograde labeling of PhMNs via phrenic nerve dip in tetramethylrhodamine. After three days, mice were euthanized, perfused with 4% paraformaldehyde, and the spinal cord excised and processed for confocal imaging. Spa mice had ~30% fewer PhMNs (P = 0.005), disproportionately affecting larger PhMNs. Additionally, a ~22% reduction in PhMN somal surface area (P = 0.019), an 18% increase in primary dendrites (P < 0.0001), and 24% decrease in dendritic surface area (P = 0.014) were observed. Thus, there are fewer larger PhMNs in spa mice. Fewer and smaller PhMNs may contribute to impaired diaphragm neuromotor control and contribute to respiratory morbidity and mortality in conditions of early onset hypertonia.

NEW & NOTEWORTHY Phrenic motor neuron (PhMN) development in early-onset hypertonia is poorly understood. Yet, respiratory disorders are a common cause of morbidity and mortality. In spa mice, an animal model of early-onset hypertonia, we found ~30% fewer PhMNs, compared with controls. This PhMN loss disproportionately affected larger PhMNs. Thus, the number and heterogeneity of the PhMN pool are decreased in spa mice, likely contributing to the hypertonia, impaired neuromotor control, and respiratory disorders.

INTRODUCTION

Motor neuron development in conditions of early-onset hypertonia, such as cerebral palsy (CP), is poorly understood. CP is the most common motor disability in children and is well recognized by the impairments in neuromotor control, especially the differences in walking ability (Palisano et al. 1997; Rosenbaum et al. 2007; Yeargin-Allsopp et al. 2008). These differences in neuromotor control affect not only limb muscles, but also respiratory muscles, with respiratory disorders being among the leading causes of death, not difficulty walking (Kwon and Lee 2013, 2014). There is a remarkable amount of research on walking differences in individuals with CP, yet very little is known about the (dys)function of the diaphragm muscle and diaphragm motor units in this population, despite significant morbidity and mortality associated with respiratory disorders.

Motor neurons are the final common output of motor control, but they are poorly understood in the context of CP (Brandenburg et al. 2019). Moreover, motor neurons are the target of many clinical therapeutic treatments for hypertonia in CP, with the goal of reducing the excitability of the motor neuron (McLaughlin et al. 2002; Quality Standards Subcommittee of the American Academy of Neurology and the Practice Committee of the Child Neurology Society 2010). Indeed, the late embryonic and postnatal periods are not the only time periods when developmental disruptions occur that may impact the brain, but are also critical time periods for spinal cord neuromotor development (Harris and McCaig 1984; Sheard et al. 1984). We have focused on the motor neuron and the spinal cord as being central to the pathogenesis of symptoms in CP. The spinal cord and motor neurons are often overlooked in investigations of CP etiology, with research focused on brain perinatal injury (Brandenburg et al. 2019), despite brain imaging often being poorly predictive for CP and severity of CP symptoms (Bax et al. 2006; Woodward et al. 2006).

To develop an effective neuromotor system, a process of programmed motor neuron death occurs during the embryonic and perinatal periods. By the start of the third trimester, the motor neuron pool reaches its maximum number before motor neuron loss occurs continuing into the perinatal period (Harris and McCaig 1984; Sheard et al. 1984). In previous studies in mice, it has been shown that the loss of effective neurotransmitter signaling for glycine (Gly) alone (Gephyrin mice), Gly and GABA (VGAT mice), or GABA alone (GAD67 mice) results in embryonic loss of putative PhMNs in the cervical spinal cord (Banks et al. 2005; Fogarty et al. 2013b, 2015). Unfortunately, all of these mouse models die shortly after birth; thus, phenotypic characterization (i.e., identifying if they develop hypertonic symptoms) and longitudinal functional studies were not possible (Banks et al. 2005; Fogarty et al. 2013b, 2015).

Orderly recruitment of motor units is essential for effective neuromotor control. It is well characterized in motor units (Henneman 1957; Henneman et al. 1965a, 1965b), including diaphragm motor units (Dick et al. 1987; Jodkowski et al. 1987), that smaller motor neurons (low capacitance, high intrinsic excitability) are recruited before larger motor neurons (higher capacitance, low intrinsic excitability). In the diaphragm, early recruited motor units accomplish ventilatory behaviors (e.g., eupnoea), and the later recruited units required to perform near-maximal expulsive/straining behaviors (e.g., coughing) (Fogarty and Sieck 2019; Mantilla et al. 2010; Sieck and Fournier 1989). It has been shown that altered glycinergic neurotransmission affects motor neuron size (Brandenburg et al. 2018; Fogarty et al. 2016, 2017a), a phenomenon which may underlie the disordered activity patterns and spasticity observed in CP.

In the spa transgenic mouse [B6.Cg-Glrb^spa/J (Gly receptor mutation)] that has reduced glycinergic neurotransmission to motor neurons (Graham et al. 2006), we have shown that the number of tibialis anterior motor neurons are markedly reduced, with a disproportionate loss of larger motor neurons (Brandenburg et al. 2018). Furthermore, we found that the number of primary dendrites on the motor neurons was increased in spa mice (Brandenburg et al. 2018). Importantly, these changes occur in the presence of severe symptoms of hypertonicity and motor impairment (Becker et al. 1986; Heller and Hallett 1982; Simon 1997). The timing, symptoms and smaller size of spa mice is homologous to many clinical features in humans with CP.

We hypothesize that reduced glycinergic neurotransmission to PhMNs during development will result in fewer numbers, reduced surface area, and an increased number primary dendrites in PhMNs of spa mice, similar to previous reports in tibialis anterior motor neurons (Brandenburg et al. 2018).

MATERIALS AND METHODS

Experimental animals and anesthesia.

This study used 10 wild-type and 5 homozygous spa knockout mice (B6.Cg-Glrb^spa/J), aged 3 to 5 mo, of both sexes. At this age, previous rodent studies have shown that glycinergic neurotransmission is mature (Becker et al. 1992; Singer et al. 1998). The B6.Cg-Glrb^spa/J knockout mice have a homozygous insertion of LINE-1 in Glrb, the β-subunit of the glycine receptor gene, resulting in a splicing error of this subunit (White and Heller 1982). Mice homozygous for this knockout (spa mice) have symptoms by the 3rd to 5th week of life, including abnormal gait, muscle rigidity, myoclonic jerks, exaggerated startle response, lower body weight, and spasticity (Becker et al. 1986; Brandenburg et al. 2018; Chai 1961; Heller and Hallett 1982; Molon et al. 2006). Founder heterozygous mice were obtained from Jackson Laboratories (Bar Harbor, ME) and used in a heterozygote × heterozygote breeding scheme. All mice used in this study were the product of this breeding scheme. All mice were housed in identical conditions. Genotyping was done on tail snips, as previously described (Brandenburg et al. 2018; Graham et al. 2006). All procedures were approved by the Institutional Animal Care and Use Committee at Mayo Clinic (Protocol no. A00003598). For survival surgery, animals received carprofen (5 mg/kg) in their water bottle starting at least 48 h before surgery for analgesia. Animals were anesthetized with intraperitoneal injections of diazepam (5 mg/kg) and droperidol (15 mg/kg) for survival surgery. Animals also received an intraperitoneal injection of fentanyl (0.3 mg/kg) and subcutaneous injection of buprenorphine SR (0.5 to 1.0 mg/kg) for additional analgesia. For terminal surgery, animals received an intraperitoneal injection of ketamine (100 mg/kg) and xylazine (10 mg/kg).

Retrograde labeling of PhMNs.

All procedures were performed using an aseptic technique, and insulated heating pads maintained body temperature at 38°C. In anesthetized mice, the phrenic nerve was identified in the ventral aspect of the neck, and the fascia was then incised and dissected to expose as much of the phrenic nerve as possible. The phrenic nerve was then cut as far distal as possible, and the proximal end of the nerve was placed in a sterile microdish with 1–3 μL of a 5% solution of tetramethylrhodamine (rhodamine) (Molecular Probes, Life Technologies, Grand Island, NY), previously validated for the accurate labeling of PhMNs (Fogarty et al. 2018b; Mantilla et al. 2009; Prakash et al. 2000). Petroleum jelly was placed on the surrounding tissues to prevent exposure to errant rhodamine solution. The retrograde labeling procedure was performed for 45 min, during which the microdish was checked at 5-min intervals, with rhodamine solution added as required (up to 9 μl total). Following this, the phrenic nerve end was removed from the microdish, and the surgical site was irrigated, cleansed, and sutured. A postsurgery period of 3 days provides ample time for retrograde transport to MNs (Gransee et al. 2013; Mantilla et al. 2009; Novikova et al. 1997; Ogilvy and Borges 1988; Zhan et al. 1989).

Confocal microscopy.

At 3 days following the retrograde PhMN-labeling procedure, animals were anesthetized and euthanized by exsanguination. Following transcardial perfusion with phosphate-buffered saline (PBS; pH 7.4) and 4% paraformaldehyde in PBS, the cervical spinal cord was postfixed in 4% paraformaldehyde overnight, and then transferred to 24% sucrose in PBS for 72 h or until sunk, before flash-freezing and cryosectioning. A cryostat (Leica Biosystems, Buffalo Grove, IL) was used to cut the samples into 70-µm longitudinal (horizontal) sections. Sections were placed on gelatin-coated slides, treated with graded ethanols and xylenes, and coverslipped with DPX mounting media (Fluka, Sigma-Aldrich, St. Louis, MO) in a manner identical to past reports (Brandenburg et al. 2018; Gransee et al. 2013, 2017; Mantilla et al. 2009).

PhMN number, primary dendrite number, and somal and dendritic surface area measurements.

Cervical spinal cord sections were imaged using an Olympus FluoView 1200 laser-scanning confocal microscope (Olympus America, Melville, NY) mounted on an upright Olympus BX50WI microscope. Rhodamine-labeled PhMNs were imaged with a ×40 oil immersion lens (NA 1.35), and three-dimensional image stacks were collected in a 1024 × 1024 array (pixel dimensions 0.50 µm × 0.50 µm) with a 2.0-µm step size. Laser intensity was 3.0 to 9.0% with confocal aperture and photomultiplier gain kept fixed across samples. Optical slices containing the nucleus of a rhodamine-labeled PhMN were identified and used to quantify the number of PhMNs. PhMN somal volumes and surface areas were determined from confocal image stacks using ImageJ (Schneider et al. 2012). Similar to past reports, we employed a stereological sampling procedure to select motor neurons for morphometric analysis (Brandenburg et al. 2018; Fogarty et al. 2018b; Prakash et al. 1994). In this procedure, PhMNs were systematically sampled in the rostral to caudal direction, and the long and short somal axis from every third PhMN was used to calculate somal surface area measurements, in a manner previously reported for prolate spheroids (Prakash et al. 1993, 1994; Ulfhake and Cullheim 1988). To determine the number of primary dendrites, systematic sampling of PhMNs in the rostral to caudal direction was performed, such that the number of primary dendrites was determined for every fifth PhMN per mouse across wild-type and spa genotypes. To determine the dendritic surface area of a particular PhMN, we measured the dendritic diameters of every 15th PhMN, in a manner identical to previous studies (Fogarty et al. 2018b; Mantilla et al. 2018). These measures were used to estimate the total dendritic surface area of a PhMN, based on a quadratic formula previously validated for rat PhMNs (Mantilla et al. 2018; Obregon et al. 2009).

Statistical analysis.

All statistical analyses were performed using standard software (Prism 7.0, GraphPad, La Jolla, CA). With respect to continuous variables, differences between groups were examined using unpaired t-tests when data were normally distributed according to D’Agostino and Pearson normality tests. A Mann-Whitney U test was used when the data were not normally distributed. Kolmogorov-Smirnov tests were used when comparing distributions, and two-way ANOVA was used when comparing two factors, with Bonferroni post hoc tests where appropriate. Linear regressions were performed with Pearson’s coefficients with a runs test to ensure no departure from linearity. Statistical significance was established at the P < 0.05 level. All experimental data are presented as means ± 95% CI, unless otherwise specified. The investigators performing the counts and morphometric measurements were blinded to the genotype of the mouse.

RESULTS

Mice characteristics.

The wild-type and spa mice were similar with respect to age and sex. However, spa mice body weights (means ± SE: 19.0 ± 3.7 g, range: 16.0–23.0 g) were 27.2% lower compared with controls (mean: 26.1 ± 3.7 g, range: 21.0–34.1 g) (Mann-Whitney U test, P = 0.018). This difference was expected on the basis of previous work with spa mice (Brandenburg et al. 2018; Molon et al. 2006). All spa mice displayed the previously reported spastic phenotype, including exaggerated startle response when handled, abnormal gait, muscle rigidity, and myoclonic jerks (Becker et al. 1986; Brandenburg et al. 2018; Chai 1961; Heller and Hallett 1982; Molon et al. 2006).

Number of PhMNs in spa mice was reduced compared with controls.

Confocal microscopy of longitudinal sections from the cervical spinal cord confirmed robust PhMN staining (Fig. 1, A and B) in both wild-type and spa mice. In spa mice, there were ~30% fewer PhMNs compared with wild-type controls (Mann-Whitney U test; P = 0.005). Wild-type mice had a mean of 211.5 ± 22.9 (range: 175–259) PhMNs, with spa mice having a mean of 147.0 ± 43.8 (range: 106–188) PhMNs (Fig. 1C). No changes in the segmental distribution of PhMNs were observed between wild-type and spa mice within the spinal cord (Fig. 1, A and B).

Fig. 1.

Reduced phrenic motor neurons (PhMNs; Phrenic MNs) in spa mice compared with wild-type controls. A and B: representative labeling of PhMNs of wild-type and spa mice, respectively. C: plots (means ± 95% CI) comparing numbers of PhMNs in control and spa mice with circles and squares representing individual animals. There was a ~30% reduction in PhMNs of spa mice (gray squares; n = 5), compared with wild-type (open circles; n = 10). Mann-Whitney U test, *P = 0.005. Scale bar: 100 µm.

Somal surface areas of PhMNs in spa mice was reduced compared with controls.

Using systematic random sampling (stereological) methodology, PhMN somal surface area measurements were obtained from 704 wild-type and 242 spa PhMNs (35–86 PhMNs per animal). Mean somal surface area was reduced by ~22% in spa mice compared with wild-type controls (wild-type: 2,261 ± 307 µm²; spa: 1,769 ± 292 µm²; Mann-Whitney U test, P = 0.019, Fig. 2).

Fig. 2.

Reduced phrenic motor neurons (PhMN; Phrenic MN) surface area in spa mice compared with wild-type controls. A and B: high-powered representative cluster of wild-type and spa PhMNs, respectively. C: plots (mean ± 95% CI) comparing PhMN somal surface areas in control and spa mice with circles and squares representing individual animals. There was a ~22% reduction in PhMN somal surface areas of spa mice (gray squares; n = 5), compared with wild-type (open circles; n = 10). Mann-Whitney U test, *P = 0.019. Scale bar: 30 µm.

Altered distribution of somal surface area of PhMNs in spa mice compared with controls.

The somal surface areas of PhMNs were calculated and plotted as frequency distributions. This frequency distribution was shifted leftward in spa mice compared with wild-type controls, indicating that the majority of PhMNs in spa mice were smaller in size (Kolmogorov-Smirnov test, P < 0.0001; Fig. 3).

Fig. 3.

Altered distribution of phrenic motor neurons (PhMN) somal surface area in spa mice compared with wild-type controls. A frequency histogram is shown (means ± 95% CI) of the number of PhMNs binned with respect to somal surface area, showing a difference in the distribution between wild-type (light gray bars) and spa (dark gray bars) mice (Kolmogorov-Smirnov test, P < 0.0001).

Fewer larger PhMNs in spa mice.

To further assess the interaction between PhMN developmental loss and the reduction in PhMN size in spa mice, PhMNs were stratified into tertiles, based on somal surface area values of wild-type controls (Fig. 4), similar to previous reports in tibialis anterior and PhMNs (Brandenburg et al. 2018; Fogarty et al. 2018b). Genotype (F_1,13 = 13.4; P = 0.003) and somal surface area tertile (F_2,26 = 4.9; P = 0.016) both had significant effects on the number of PhMNs. For spa mice, there was a significant reduction in the number of PhMNs in the upper tertile compared with lower tertile (P = 0.011) (Fig. 4). However, there was no difference between the middle and lower tertiles (P = 0.326) nor between the middle and upper tertiles (P = 0.409) (Fig. 4). In comparison to all tertiles, in wild-type mice, the number of PhMNs in the upper tertile of spa mice was reduced (P < 0.044) (Fig. 4).

Fig. 4.

Disproportionate loss of larger phrenic motor neurons (PhMNs) in spa mice compared with wild-type controls. Histogram of the estimated number of PhMNs (means ± 95% CI) within wild-type (open bars; n = 10) and spa (gray bars; n = 5) mice, stratified into lower, middle, and upper tertiles, based on the distribution of PhMN somal surface area of wild-type controls. Hence, there are no differences between tertiles in wild-type mice. In spa mice, the number of PhMNs in the upper tertile was significantly reduced, as compared with all wild-type tertiles (P < 0.05). In addition, upper tertiles spa PhMNs were significantly reduced compared with lower tertile PhMNS (P < 0.05). Thus, spa mice have a disproportionate loss of the larger, upper tertile PhMNs. Two-way ANOVA with post hoc Bonferroni testing; letters that differ denote P < 0.05.

Number of primary dendrites at PhMNs in spa mice was increased compared with controls.

Using systematic random sampling (stereological) methodology, PhMN primary dendrite numbers were obtained from 424 wild-type and 152 spa PhMNs (21–52 PhMNs per animal). The mean number of primary dendrites for PhMNs in spa mice was ~18% greater as compared with wild-type controls (wild-type: 4.0 ± 0.1; spa: 4.9 ± 0.3; unpaired t-test, P < 0.0001, data not shown).

Reduced total dendritic surface areas of spa PhMNs compared with controls.

Using systematic random sampling (stereological) methodology, PhMN primary dendrite diameters were obtained, and total dendritic surface areas were estimated from every 15th PhMN from both wild-type and spa mice. The mean total dendritic surface area of PhMNs in spa mice (16,410 ± 5,940 µm²) was ~24% less as compared with wild-type controls (21,730 ± 1,838 µm²; unpaired t test, P = 0.014, Fig. 5A).

Fig. 5.

Disproportionate reduction of total dendritic surface area of larger spa phrenic motor neurons (PhMNs) compared with wild-type controls. A: a plot of the estimated total dendritic surface area of PhMNs is shown [mean ± 95% confidence interval (CI)], with ~24% reduction in spa (n = 5) total dendritic surface area compared with wild-type (n = 10) mice (P = 0.014, unpaired t-test). B: histogram of the estimated total dendritic surface area of PhMNs (mean ± 95% CI) in wild-type (open bars, n = 10) and spa (gray bars, n = 5) mice, stratified into lower, middle and upper tertiles, based on wild-type PhMN somal surface area distributions. The total dendritic surface area of PhMNs in the upper tertile in spa mice was significantly reduced compared with wild-type mice (*P < 0.05, two-way ANOVA with Bonferroni post hoc tests). C: linear relationship between PhMN somal surface area and estimated total dendritic surface areas in both wild-type (slope = 7.1, P < 0.0001; r² = 0.55) and spa mice (slope = 4.7, P < 0.0001; r² = 0.32). The slope of this linear relationship is reduced by 34% in spa mice, indicative of reduced dendritic surface areas in larger spa PhMNs (P = 0.01).

To assess the interaction between PhMN size and total dendritic surface area, PhMNs were stratified into tertiles based on somal surface area values of wild-type controls (see Fig. 4B). The mean values of total dendritic surface area were calculated for each mouse within each somal surface area tertile. Genotype (F_1,13 = 16.1; P = 0.002), somal surface area tertile (F_2,22 = 40.8; P < 0.0001), and the interaction between both (F_2,26 = 9.8; P = 0.001) had significant effects on the total dendritic surface area of PhMNs. For spa mice, there was a 34% reduction in the total dendritic surface area of PhMNs in the upper tertile compared with wild-type (P = 0.02) (Fig. 5C). There was no difference in total dendritic surface area between spa and wild-type mice of the lower (P > 0.99) or middle (P = 0.12) size tertiles (Fig. 5B).

We also plotted each individual PhMN total dendritic surface area against the somal surface area for both spa and wild-type mice. Similar to past reports evaluating dendritic length (Fogarty et al. 2019), wild-type PhMNs exhibit a strong linear correlation between somal surface area and total dendritic surface area (slope = 7.1, P < 0.0001; r² = 0.55; Fig. 5C). A linear correlation was also evident in spa mice (slope = 4.7, P < 0.0001; r² = 0.32), although there was a significant difference (34%) in the slope of the regression lines between spa and wild-type (F_1,227 = 6.6, P = 0.01; Fig. 5) mice.

DISCUSSION

This study had four main findings: 1) spa mice have fewer PhMNs than wild-type mice, 2) PhMNs in spa mice are smaller, 3) there is a disproportionate loss of larger PhMNs in spa mice, 4) the total dendritic surface areas of PhMNs in spa mice are reduced, despite having a greater number of primary dendrites, and 5) dendritic surface area reductions disproportionately affect larger PhMNs in spa mice.

Altered glycinergic neurotransmitter input to PhMNs, via a Gly receptor mutation results in increased developmental loss and fewer PhMNs in spa mice compared with wild-type mice. It has been consistently demonstrated that fetal motor neuron developmental loss is affected by alterations of glycinergic and GABAergic signaling with an ~30% loss of motor neurons in cervical segments involved in innervating the diaphragm of fetal gephyrin, VGAT, and GAD67 mice (Banks et al. 2005; Fogarty et al. 2013b, 2015). However, these previous studies did not unambiguously identify PhMNs by retrograde labeling. In our present study, we found an ~30% reduction in the number of identified PhMNs in adult spa mice, similar to observations of putative PhMNs in gephyrin, VGAT, and GAD67 mutants. This suggests that the loss of PhMNs occurs at some point in development, as opposed to later in adulthood. PhMN loss is likely related to rostral caudal timing in transition of GABA and Gly from excitatory to inhibitory neurotransmitters and the relative abundance of GABA and Gly neurotransmission. In PhMNs of healthy rodents, the postnatal transition to inhibitory action occurs by ~2 wk after birth, a time when developmental loss of motor neurons is complete (Ben-Ari 2002; Kriegstein and Owens 2001; Wong-Riley and Liu 2008) and before the emergence and expansion of DIAm fibers, expressing MyHC_2X and MyHC_2B isoforms (Mantilla et al. 2017; Prakash et al. 2000). In spa mice, studies in postnatal day (P) ~25 mice show that Gly has inhibitory action on hypoglossal motor neurons (Graham et al. 2006; Tadros et al. 2014) and in spinal cord dorsal horn interneurons (Graham et al. 2003). Our animals were between P90 and 150, being well beyond when effective GABA and Gly neurotransmission has transitioned to inhibitory influence in the spinal cord. Loss of effective signaling of either GABA or Gly neurotransmission would result in a decrease in excitatory input to motor neurons early in development and a loss of inhibitory input (i.e., disinhibition) in adults.

In our study, the location of PhMNs was concentrated at the C₃, C₄, C₅, and C₆ spinal cord segments, consistent with previous reports in mice and rats (Goshgarian and Rafols 1981; Vandeweerd et al. 2018; Qiu et al. 2010). However, the absolute number of PhMNs in wild-type mice in our study differs somewhat from previous reports. Our PhMN counts greater than the number of PhMNs reported in adult mice using either cholera toxin B painting on the diaphragm (~190 counts) (Qiu et al. 2010) or intrapleural cholera toxin b injection (~150–190 counts) (Vandeweerd et al. 2018). The differences in PhMN numbers are most likely due to differences in technique, with nerve dip labeling more PhMNs than intrapleural or intramuscular injections of cholera toxin B (Mantilla et al. 2009). In previous studies in rats, we used phrenic nerve dip as the “gold standard” for assessing PhMN number (Fogarty et al. 2018b; Mantilla et al. 2009; Prakash et al. 1994, 2000). To ensure complete PhMN labeling, we employed the nerve dip technique, avoiding any confounding factors due to potential abnormalities in spa phrenic axon terminals that would impair the uptake of intrapleural or intramuscular cholera toxin B (reliant on uptake via gangliosides on the nerve terminals).

In addition to fewer PhMNs in adult spa mice, we found that PhMNs in spa mice had smaller somal surface areas. The shift in the distribution of somal surface areas in spa mice could be attributed to a disproportionate loss of larger PhMNs or a decrease in the size of surviving PhMNs in spa mice. However, the number of PhMNs in the lower tertiles was preserved in spa mice compared with wild-type, suggesting a selective loss of larger PhMNs. This has implications on functional diaphragm behaviors as larger PhMNs innervate more fatigable fast-twitch motor units (type FInt and FF motor units), which consist of type IIx and/or IIb muscle fibers (Fournier and Sieck 1988; Mantilla and Sieck 2008; Sieck et al. 1989, 1996). These FInt and FF motor units develop only after ~2–3 wk in rodents (Mantilla et al. 2008), with the appearance of type IIx and/or IIb muscle fibers. FInt and FF motor units are responsible for generating higher-force diaphragm behaviors, which are critical in airway clearance (i.e., coughing) and Valsalva (i.e., defecation) maneuvers (Fogarty et al. 2018a; Fogarty and Sieck 2019). Respiratory disorders are a leading cause of death in individuals with CP, and although the etiologies of the respiratory disorders are likely to be multifactorial, impairments in the ability to clear the airway is likely a significant factor. Constipation is also a common condition that affects a majority of individuals with CP, and again, while constipation is likely multifactorial, difficulty eliciting high-force diaphragm contraction to increase intraabdominal pressure and facilitate expulsion of stool may be a significant factor. In the spa mouse, it is not clear whether there is a loss of larger PhMNs or whether PhMNs fail to grow to a large size during postnatal development (Prakash et al. 2000). Overall, similar to our previous work on lumbar spinal cord motor neurons innervating the tibialis anterior muscle (Brandenburg et al. 2018), our results in spa mice suggest that there is a selective vulnerability of larger PhMNs that comprise FInt and FF motor units.

The reduced somal surface areas of both tibialis anterior motor neurons and PhMNs in spa mice is consistent with increased intrinsic motor neuron excitability, which corresponds with the hypertonic phenotype. According to the Henneman Size Principle, motor neurons with smaller cell bodies are recruited before those with larger cell bodies reflecting greater intrinsic excitability (Henneman 1957; Henneman et al. 1965a, 1965b). Furthermore, this intrinsic excitability may not be the sole contributor, as we recently demonstrated that smaller PhMNs have a greater density of glutamatergic inputs (i.e., excitatory inputs) than larger PhMNs (Rana et al. 2019). In addition to smaller PhMN somal size and possibly a greater density of glutamatergic inputs to PhMNs in mice, the reduced influence of inhibitory Gly input onto PhMNs would contribute to hyperexcitability and hypertonia.

In the present study, we observed that PhMNs have more primary dendrites in spa mice as compared with wild-type mice. Although spa PhMNs had more dendritic trees, their total dendritic surface areas were reduced, particularly in larger PhMNs. This result is consistent with a developmental restriction in the growth of the spa PhMNs that would comprise FInt and FF motor units. The technique of retrograde rhodamine labeling of PhMNs used in the present study reliably identifies dendrites up to the 3rd dendritic branch (Fogarty et al. 2018b; Issa et al. 2010; Prakash et al. 2000) and cannot assess the full dendritic tree of PhMNs, which can extend to the 8th order (Fogarty et al. 2017b, 2019; Leroy et al. 2014). Previous studies of inhibitory neurotransmission defects (including glycine receptor mutations) in motor neurons also support the concept that an imbalance of excitation/inhibition inputs can alter dendritic trees in motor neurons (Fogarty et al. 2016, 2017a). Future studies could make use of alternative techniques that permit detailed evaluation of the dendritic tree, including Golgi-Cox staining (Fogarty et al. 2017b, 2019), or intracellular labeling (Fogarty et al. 2013a; Kanjhan et al. 2016; Obregon et al. 2009; Pace et al. 2002).

Although loss of a single neurotransmitter input is not a known etiology for CP, spa mice display a well-described hypertonic phenotype with abnormal locomotion, muscle rigidity, myoclonic jerks, and an exaggerated startle response along with a lower body weight (Becker et al. 1986; Brandenburg et al. 2018; Heller and Hallett 1982). Thus, the spa mice phenotype has remarkable similarities to individuals with CP. As the phenotype is critical to the diagnosis of CP, animals with a CP-like phenotype are critical to furthering the understanding of mechanisms underlying this condition (Brandenburg et al. 2019). Because of their phenotype, spa mice have precedence for modeling CP in studies exploring clinical findings and treatments in CP (Cosgrove and Graham 1994; Ziv et al. 1984). Furthermore, as spinal cord development lags brain development (Eyre et al. 2000), any injury that affects a developing brain will also affect development of the spinal cord and the motor neurons. Thus, hypertonia in humans with CP is likely not only due to an early brain injury causing an imbalance of excitatory and inhibitory neurotransmitters, but also a developmental disruption of motor neurons in the spinal cord. Specifically, disruption of Gly, which is expressed in the spinal cord (van den Pol and Gorcs 1988; Werman et al. 1967, 1968), appears to have a key role in the abnormal PhMN loss and development of hypertonia, similar to what we have previously shown with tibialis anterior motor neurons in the lumbar spinal cord (Brandenburg et al. 2018). As spa mice survive into adulthood, this mouse model provides a unique opportunity to longitudinally determine the impact of impaired neurotransmitter signaling on PhMN, diaphragm motor unit, and diaphragm muscle mechanical properties throughout the entire lifespan. Evaluating PhMN properties in mature spa mice is the first step toward this goal.

GRANTS

This study was supported by the National Institutes of Health (NIH) Grants R01-AG-044615 (to G. C. S.) and R01-HL-96750 (to G.C.S.), an Australian National Health & Medical Research Council C. J. Martin Early Career Fellowship (to M.J.F.), Mayo Clinic Office of Research Diversity and Inclusion Career Support and Advancement Award (to J.E.B.), and Richard and Rosemary Crandall (to J.E.B.).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

J.E.B., M.J.F., and G.C.S. conceived and designed research; J.E.B., M.J.F., and A.D.B. performed experiments; J.E.B. and M.J.F. analyzed data; J.E.B., M.J.F., and G.C.S. interpreted results of experiments; J.E.B. and M.J.F. prepared figures; J.E.B. drafted manuscript; J.E.B., M.J.F., A.D.B., and G.C.S. edited and revised manuscript; J.E.B., M.J.F., A.D.B. and G.C.S. approved final version of manuscript.