Blunted Respiratory Responses in the Streptozotocin-Induced Alzheimer’s Disease Rat Model

Abstract

Alzheimer’s disease (AD) is known for the progressive decline of cognition and memory. In addition to these disease-defining symptoms, impairment of respiratory function is frequently observed and often expressed by sleep-disordered breathing or reduced ability to adjust respiration when oxygen demand is elevated. The mechanisms for this are widely unknown. Postmortem analysis from the brainstem of AD patients reveals pathological alterations, including in nuclei responsible for respiratory control. In this study, we analyzed respiratory responses and morphological changes in brainstem nuclei following intracerebroventricular (ICV) injections of streptozotocin (STZ), a rat model commonly used to mimic sporadic AD. ICV-STZ induced significant astrogliosis in the commissural part of the nucleus tractus solitarii, an area highly involved in respiration control. The astrogliosis was identified by a significant increase in S100B-immunofluorescence that is similar to the astrogliosis found in the CA1 region of the hippocampus. Using plethysmography, the control group displayed a typical age-dependent decrease of ventilation that was absent in the STZ rat group. This is indicative of elevated minute ventilation at rest after STZ treatment. Peripheral chemoreflex responses were significantly blunted in STZ rats as seen by a reduced respiratory rate and minute ventilation to hypoxia. Central chemoreflex responses to hypercapnia, on the other hand, only decreased in respiratory rate following STZ treatment. Overall, our results show that ICV-STZ induces respiratory dysfunction at rest and in response to hypoxia. This provides a new tool to study the underlying mechanisms of breathing disorders in clinical AD.

Keywords

Astrocytosis brainstem chemoreflex hypercapnia hypoxia nucleus tractus solitarii ventilation

INTRODUCTION

Alzheimer’s disease (AD) has debilitating effects on memory and cognition [1]. An accompanying feature of the disease are multiple forms of autonomic and respiratory dysfunction [2 –4]. Respiratory deficits are prevalent in AD patients and often manifest in sleep-disordered breathing (SDB) [5]. A common form of SDB is obstructive sleep apnea with its recurring episodes of intermittent hypoxia. Sleep apnea is associated with an increased risk of dementia [6], and markers of AD are found in children with obstructive sleep apnea [7]. Furthermore, there is a strong correlation of the severity of SDB with the severity of AD [8]. Challenging the respiratory system of AD patients showed a reduced ability to increase respiration, as seen by blunted minute ventilation to a heightened oxygen demand with peak efforts in exercise [3]. Also, late stages of clinical AD are accompanied by shortness of breath [9].

The underlying mechanisms for these alterations in the course of AD are unknown. It was shown that clinical AD also impacts regions in the lower brainstem, including those important for respiratory control [10, 11]. To unravel the respiratory changes in AD, it is important to find suitable animal models that display this aspect of the disease. Thus, we turned toward the streptozotocin (STZ)-induced rat model of AD that closely matches the multifactorial pathogenesis and symptoms of sporadic AD (95% of AD cases in humans) [12 –22]. With intracerebroventricular injections of STZ, we detected astrogliosis in the commissural nucleus tractus solitarii (nTS), a brainstem area integral to the control of respiration. Analysis of respiratory parameters using plethysmography revealed increased minute ventilation at rest and significantly blunted chemoreflex responses to hypoxia.

MATERIALS AND METHODS

Animals

Male Sprague-Dawley rats (Harlan; n = 28, 272±8 g) were maintained in the AAALAC accredited vivarium of the ATSU Kirksville College of Osteopathic Medicine. Rats were held on a 12-h day/night cycle at 24°C and 46% humidity with water and food available ad libitum. A subset of experiments was conducted at the University of Missouri and rats were held under the same conditions in the AAALAC accredited vivarium of the Dalton Cardiovascular Research Center. All experimental protocols are in accordance with NIH guidelines (“Guide for the Care and Use of Laboratory Animals”) and approved by the Animal Care and Use Committee of A.T. Still University and University of Missouri.

Intracerebroventricular injection of streptozotocin

Similar to previous studies [20, 23], STZ was injected into both lateral ventricles of the brain. We used a sub-diabetogenic dose of STZ [24, 25] as indicated by peripheral blood glucose levels that were comparable to control rats (CTL) injected with artificial cerebrospinal fluid (aCSF/vehicle, n = 6, 85.5±5.3 mg/dL blood versus STZ, n = 5, 98.8±10.7 mg/dL blood; p = 0.27). Blood glucose levels were tested immediately before euthanasia using a commercially available blood glucose tester (TRUEresult, Nipro diagnostics). For ICV injections, rats were anesthetized using isoflurane (Piramal; 5% induction, 2% maintenance) and positioned in a stereotaxic frame (David Kopf Instruments). Before injections, rats received 2 mg/kg Dexamethasone (Med-Pharmex; immunosuppressant) to prevent possible brain swelling. A midline sagittal incision was made in the scalp to expose the coronal and sagittal suture of the skull. Two burr holes were drilled into the skull with a rotary tool (Dremel 7300 with engraving cutter 105) to allow access to the lateral ventricles. Injection glass capillaries (World Precision Instruments) were pulled using a P-97 micropipette puller (Sutter Instruments) and used for ICV-injections according to the following stereotaxic coordinates: –0.9 mm AP,±1.5 mm ML, and 3.6 mm DV [26]. Figure 1 shows the typical injection site and injection canal through the cortex marked with 2% methylene blue. 25 mg/mL STZ was dissolved in aCSF (in mM: 123 NaCl, 1.1 CaCl₂, 3 KCl, 1.9 MgCl₂, 25 NaHCO₃, 0.5 NaH₂PO₄, and 0.25 Na₂HPO₄, pH 7.3) and pressure-injected at a variable volume (<10 μL per side) to achieve a dose of 1.5 mg STZ per kg rat. Control animals received vehicle only. The wound was sutured shut using absorbable suture (4-0, Oasis, MV-J397-V) and rats were postoperatively treated with 50 μg/kg Buprenorphine hydrochloride (Sigma) for pain management, 7 mg/kg Baytril (Bayer) for antibiotic treatment, and 3 mL 0.9% sodium chloride solution (Hospira) for fluid reconstitution. The surgical procedure for a second ICV injection was repeated 2 days later. Within the first week following surgery, rats were supplemented with sweetened cereal (Froot loops, Kellogg’s) for weight recovery.

Passive avoidance memory test

Similar to previous studies [27, 28], we validated our model to confirm memory decline in a subset of rats using the passive avoidance step-through paradigm. The avoidance system (GEMINI avoidance system, San Diego Instruments) consisted of a light and a dark chamber separated by a guillotine door. In the acquisition trial 14 days following ICV injections, rats were placed in the light chamber. Following 2 min of habituation, the guillotine door was raised and the time recorded until the rat entered the dark chamber. There was no difference in initial crossing latency between CTL and STZ rats (CTL, n = 5, 62±21 s versus STZ, n = 3, 34±10 s; p = 0.786). Immediately after entering the dark chamber, rats received a 3-s electric foot shock (0.4 mA) though the metal floor grids and were then transferred back to their cage. In the retention trial 24 h later, rats were subjected to the same procedure (without the electric foot shock). All CTL rats remained in the light chamber for more than 300 s (cut-off time). On the other hand, all STZ rats showed impaired recall latency as indicated by a significant shorter crossing latency into the dark chamber (CTL, n = 5, 300±0 s versus STZ, n = 3, 126±77 s; p = 0.036).

Immunohistochemistry

Similar to our previous manuscripts [29, 30], immunohistochemistry was performed in a subset of rats two weeks following the second ICV injection. Rats were deeply anaesthetized with isoflurane and transcardially perfused with heparinized 0.01 M phosphate-buffered saline (PBS; in mM: 2.7 NaH₂PO₄, 7.7 Na₂HPO₄, and 154 NaCl, pH 7.4) that was followed by 4% paraformaldehyde (PFA, Acros organics) in PBS. The brain was removed, post-fixed overnight, cut into 30 μm thick coronal sections using a vibrating microtome (7000smz-2, Campden Instruments), and collected either in a 1/6 series (brainstem; 24 well-tissue culture plate) or 1/4 series (forebrain; 12 well-plates). Sections were washed in PBS and blocked for 30 min in 10% normal donkey serum (NDS, Millipore) with 0.3% Triton-PBS. Subsequently, brain sections were incubated in 1% NDS with 0.3% Triton-PBS and the primary antibody against S100 calcium-binding protein B (S100B; astroglial marker; rabbit, 1:200, ab41548, Abcam). The following day, sections were rinsed in PBS and incubated for 2 h in 1% NDS with 0.3% Triton-PBS and the secondary antibody Alexa Fluor 488-conjugated donkey anti-rabbit IgG (1:200; 711-545-152, Jackson Immuno). Brain sections were mounted on gelatin-coated glass slides, air dried for 1 h, and cover-slipped using ProLong Diamond with 4’,6-diamidino-2-phenylindole (ThermoFisher Scientific). One section was incubated in the absence of the primary antibody and showed no fluorescent staining (negative control). While all hippocampal sections were simultaneously labeled against S100B using the same solutions, tissue sections from the brainstem were processed in pairs consisting of one aCSF control and one STZ rat. Specificity of the S100B antibody was previously confirmed [31].

Immunoreactivity was generally visualized using a conventional epifluorescent microscope (Eclipse 80i, Nikon), a digital monochrome camera (DS-Qi1Mc, Nikon), and appropriate filter sets and excitation wavelength for the fluorophore. Figure 2D is a maximum projection of 8 images taken in a z-stack (0.5 μm) with a 63x (oil) objective on a spinning disc confocal microscope (DMI6000 B, Leica) with CCD camera (ImagEM Enhanced, Hamamatsu). Exposure times were kept identical for all hippocampal section or tissue that was processed in pairs to allow comparison between both treatment groups. Images were post-processed using ImageJ (Version 1.49t, NIH) by rescaling pixel size (bicubically) and adjusting contrast and brightness (identical histogram range for all slice pair comparisons). The clarity of Fig. 2D was enhanced by deconvolution (10 iterations, blind PSF, medium noise) using AutoQuant X3 (Media Cybernetics). According to the fluorophore of the secondary antibody images were given the pseudo-color green. Templates shown in Figs. 1, 2, and 3 were drawn according to coronal brain slices using the freeware program Inkscape (version 0.91; http://www.inkscape.org/). The position from Bregma and location of some brain structures were determined using the rat atlas from Paxinos & Watson [26]. In each animal, one corresponding brain section per brain region (hippocampus, nTS, pre-Bötzinger complex) was used to analyze astrocyte number and process density. Astrocyte numbers were manually counted by two blinded individuals in a 330×330 μm box for the CA1 region of the hippocampus, a 100×100 μm box for the commissural nTS, and a 150×150 μm box for the pre-Bötzinger complex using ImageJ. Box sizes were chosen to include most of the target area while excluding potential differences in nucleus size. S100B staining of astrocytic processes was quantified as mean staining intensity in ImageJ using the original grey-scale images. Measurements were taken adjacent to the cell bodies (see Fig. 2A & D) and represent an average of multiple measurements within the nucleus of interest for each animal. In the hippocampus, each average consists of measurements using 30×30 μm boxes at three different locations. Five measurements using 10×10 μm boxes were used for the other nuclei. Staining intensity from tissue that was processed in pairs was normalized to aCSF control to allow group comparison.

Plethysmography

Prior to any surgical procedures (pre-aCSF/STZ) and 14 days following the second ICV-injection (post-aCSF/STZ), respiratory parameters were recorded in conscious unrestrained rats using a whole-body plethysmography chamber (Data Sciences International). Pressure changes from respiration of the rat inside the test chamber were compared to those in a reference chamber and measured with a differential pressure transducer (DP45, Validyne). The test chamber was continuously flushed with bias gas flow of 3 L/min, making the recorded pressure signal directly proportional to changes in air flow inside the chamber. The pressure signal was demodulated (Sine wave carrier demodulator CD15, Validyne), digitized (PowerLab 8/35, AD Instruments), and recorded via LabChart Software (AD Instruments). Rats were exposed to different gas mixtures by blending pure gases using two mass flow controllers (MC-5SLPM-D) and control software (Flow Vision MX) with hardware interface (RS-232 Multi-Drop, Model BB9) from Alicat Scientific. For the experiments conducted at the University of Missouri pure gases were mixed using the gas blender Hypoxydial (Starr Life Sciences). The other devices utilized for plethysmography were similar at both institutes. Rats were acclimated to the plethysmography chamber for 2 days (1 h each) prior experimental protocols. On the day of the experiment, gas mixtures were given (following stabilization of breathing at 21% O₂) for 7 to 10 min to allow adequate time for gas exchange within the chamber and chemoreflex responses. To stimulate the peripheral or central chemoreflex, rats were exposed to hypoxia (14, 12, 10, and 8% O₂, equilibrated with N₂; following 21% O₂) or hypercapnia (5% CO₂, equilibrated with O₂; subsequent to 20 minutes of 100% O₂ to eliminate influence of the peripheral chemoreflex), respectively. Data were analyzed using LabChart and Microsoft Excel Software. Respiratory rate (breath per minute) and tidal volume (normalized to 100 g rat) were measured during steady and uninterrupted breathing (in the absence of movement, sniffing, and sighs) within the last 30 s of a gas mixture. Minute ventilation (per 100 g rat) was calculated as the product of respiratory rate and tidal volume/100 g. Rate and volume of sighs (i.e., augmented breaths; per 100 g rat) were analyzed within the last 3 min of a given gas mixture. Sigh and tidal volume were calculated as the area under the inspiratory pressure curve and by using a formula previously described to correct for barometric pressure, temperature, and water vapor pressure inside the animal and plethysmography chamber [32]. Volume measurements were calibrated to a pressure change of a known volume of air injected into the chamber using a rodent respirator (Harvard Apparatus, Model 680).

Statistical analysis

Statistical analyses were performed using SigmaPlot 13.0 (Systat Software). Immunohistochemical data and percent changes during the central chemoreflex were analyzed using paired t-tests. Data from the passive avoidance test were compared using the Mann-Whitney rank sum test. All other data were compared by two-way repeated measures ANOVA (with Student–Newman–Keuls post hoc test). Results are considered significantly different at p values≤0.05. Group data are presented as means±SEM.

RESULTS

STZ-injections induce astrogliosis in the hippocampus and commissural nTS

Reactive astrocytes (astrogliosis) are highly associated with clinical AD [33]. An increase in astroglia has also been previously demonstrated in the hippocampus of the STZ-induced AD rat model using immunoblots [20]. To verify that similar alterations occur in this study, we immunohistochemically labeled astrocytes in the hippocampus using S100B antibody. Figure 2A shows a typical staining pattern of astrocytes in the CA1 region of the hippocampus (box in Fig. 2B) from aCSF-injected control (CTL) rats and STZ-injected rats. There was no statistical difference in the number of astrocytes between the CTL and STZ rat group (Fig. 2C). However, since astrogliosis is mainly characterized by an increased thickening and number of cellular processes [34, 35], we measured S100B staining intensity adjacent to the cell bodies. Typically, the network of astrocytic processes in the CA1 region had a higher density in STZ rats when compared to CTL (Fig. 2D). This was supported by a significant increase of S100B staining intensity in the group data (Fig. 2E).

To identify if astrogliosis also affects respiratory centers in the brainstem, we analyzed S100B immunoreactivity in the commissural nTS and pre-Bötzinger complex representatively for the dorsal and ventral respiratory group (Fig. 3). Similar to the hippocampus, the commissural nTS of STZ rats exhibited a consistently higher degree of S100B staining intensity when compared to CTL rats, while the number of astrocytes was unchanged (Fig. 3A-C).The pre-Bötzinger complex (Fig. 3D, E), again, showed no change in astrocyte number (Fig. 3F left). However, and in contrast to the commissural nTS and hippocampus, S100B staining intensity was not altered (Fig. 3F, right).

Increased resting respiration with ICV STZ

Astrogliosis within the commissural nTS may be associated with alteration of respiratory parameters. Thus, we analyzed respiratory rate, tidal volume, and minute ventilation using whole-body plethysmography in conscious rats. Baseline values (at 21% O₂) before (pre-) and two weeks following (post-) ICV injections of aCSF or STZ are given in Table 1. While most parameters remained constant within the two treatment groups, resting minute ventilation fell significantly in CTL rats over time. This decrease is similar to the reduction of ventilation with rat age when adjusted for body weight [36 –38]. STZ rats, however, did not show the time-dependent decrease in ventilation. These results indicate increased respiration at baseline in response to STZ treatment.

Peripheral chemoreflex responses are bluntedin STZ rats

To evaluate possible alterations in the peripheral chemoreflex, we analyzed respiratory parameters in response to hypoxia (14–8% O₂). Reducing environmental O₂ content within the plethysmography chamber reliably elevated respiratory rate in CTL and STZ rats (p≤0.001, n = 8 for each group). The representative example shown in Fig. 4A exhibited a maximum respiratory rate at 10% O₂ that slightly dropped with reduction of O₂ to 8%. These changes in respiratory rate are similar in both rat groups prior to any intervention (open circles/squares Fig. 4B, top). Following ICV injections, post hoc analysis revealed a significant reduction of respiratory rate in the STZ rat, whereas CTL rats did not change (closed circles/squares Fig. 4B, top). Tidal volume, on the other hand, did not change over most hypoxic severities tested (Fig. 4B, middle). At 8% O₂, however, both rat groups had significantly increased tidal volume before treatment as shown by post hoc analysis (p≤0.01, n = 8 for each group, against all other hypoxic severities). Two weeks after ICV injections this significant increase in response to hypoxia was absent in the STZ-treated group but not the control group. Minute ventilation was steadily rising with increasing hypoxia (Fig. 4B, bottom; p≤0.001, n = 8 for each group). In the STZ rat group this rise in minute ventilation was significantly blunted after STZ treatment. CTL rats were similar pre- and post-aCSF treatment.

Normalizing the data sets from Fig. 4B to their baseline values at 21% O₂ allows assessment of chemoreflex gain independent of the changes in resting ventilation over time that were observed in CTL rats. Fig. 5 shows the difference between pre- and post-injection of normalized respiratory rate, tidal volume, and minute ventilation during hypoxia for CTL and STZ rats. Even though there were no statistically significant changes between rat groups and hypoxic severities for respiratory rate and tidal volume (Fig. 5, top and middle), respiratory rate in CTL rats was generally increased at post-injection. The response of AD rats, on the other hand, was reduced by more than 20% at 14% O₂. This decrease gradually disappeared with increasing severity of hypoxia. Tidal volume remained stable during 14–10% O₂ in both rat groups. At 8% O₂, however, the response of STZ rats showed a decrease of about 30% in comparison to the pre-injection data. Minute ventilation (Fig. 5, bottom) of CTL rats was increased to each hypoxia tested. Responses of STZ rats, in contrast, were consistently decreased by more than 20% for each hypoxic severity tested. At 10% and 8% O₂ this change in minute ventilation was significantly different from control, indicating a progressive blunting of chemoreflex gain with increased severity of hypoxia.

The central chemoreflex shows little impairment in STZ rats

With the STZ-induced blunting of ventilatory response to hypoxia, central chemoreflex responses to hypercapnia may also be altered. This was tested by increasing the CO₂ concentration (5% CO₂ equilibrated with O₂) subsequent to 20 min of 100% O₂.The high concentration of O₂ was given to minimize any influence of the peripheral chemoreflex. Prior to aCSF or STZ injections, hypercapnia induced a robust increase in breathing rate, tidal volume, and minute ventilation (p≤0.01, n = 5 for each group) as shown by the raw values in Fig. 6A (open circles/squares). These responses did not change following aCSF injection. STZ rats showed a significantly reduced respiratory rate during 5% CO₂ when compared to its baseline value (i.e., before STZ injection). This decrease in respiratory rate in combination with unchanged tidal volume, however, did not alter minute ventilation. Normalizing these data to 100% O₂ (Fig. 6B) shows minimal changes in both rat groups for respiratory rate, tidal volume, and minute ventilation.

STZ rats demonstrate augmented sigh amplitude with hypoxia

Sighs are augmented breaths that occur intermittently during normal breathing to improve lung oxygenation by reopening collapsed alveoli [39]. Sigh frequency generally increases with hypoxia [40] and thus is indicative of the hypoxic state of the animal. We analyzed sigh frequency and volume in the CTL and STZ rat. Fig. 7A shows a typical example of intermittent sighs during the breathing rhythm when exposed to hypoxia. Similar to previous reports [41], sighs can be distinguished from regular breaths by its augmented amplitude and its biphasic shape (Fig. 7A, bottom). The initial phase (I) appears similar to the inspiratory part of a regular breath. In phase II, the amplitude of inspiration is greatly extended. The mean data shows that sigh frequency increases with the severity of hypoxia (Fig. 7B, top open circles/squares; p≤0.001, n = 8 for each group). Neither treatment with aCSF nor STZ changed the magnitude of this relationship (Fig. 7B, top closed circles/squares). Sigh volume, in contrast, generally decreased with hypoxic severity (Fig. 7B, bottom; p≤0.001, n = 8 for each group). Following ICV injection, the fall of sigh amplitude to hypoxia was significantly attenuated in STZ rats.

DISCUSSION

In this study, we demonstrated that ICV-STZ induces astrogliosis within the commissural nTS that is similar to the astrogliosis found in the hippocampus. Respiration of STZ rats was significantly altered as seen by increased minute ventilation at rest and a blunted peripheral chemoreflex response to hypoxia. Additionally, sigh volume of STZ rats was significantly increased during hypoxia when compared to pre-injection values. On the other hand, STZ treatment had little effect on central chemoreflex responses to CO₂. This study provides the first evidence on respiratory dysfunction and morphological alteration of an associated respiratory center with ICV-STZ.

Pathological changes with STZ injections closely match the symptoms and pathogenesis of human AD [12 , 43]. Among others, alterations with ICV STZ include decreased memory and cognition, inflammation, oxidative damage, and gliosis, as well as amyloid-β plaques and phosphorylated tau. In the present study, we validated our model by showing impaired memory and astrogliosis in the hippocampus of STZ rats. Memory impairment is a hallmark of AD and frequently observed in the STZ-induced AD model [24, 28]. Furthermore, astrogliosis is considered as an early key event of several AD models [20 , 44–46], including the STZ-induced model of sporadic AD used in this study. Astrogliosis is also highly associated with human AD [33 , 48] and its degree inversely correlates with cognitive function [49]. Finally, we also verified that ICV STZ does not alter blood glucose levels, since peripheral injections of STZ is commonly used as a model for diabetesmellitus [50].

With the morphological alterations in the hippocampus, we found astrogliosis in the nTS of the brainstem. The nTS is the initial brain center for information from multiple visceral afferents [51] and essential for autonomic and respiratory control [51 –53]. The nTS is starting to receive more attention due to its important role in the control of autonomic responses and the morphological changes observed in the course of AD [11, 54]. Thus, plastic changes within the nTS likely affect the function of the respiratory system. Apart from this, our data did not indicate astrogliosis in the pre-Bötzinger complex, which is a primary nucleus that houses many of the central pattern generators of the respiratory network [55]. The STZ-induced changes in this study may represent early events in AD (two weeks following STZ injections) and astrogliosis in the pre-Bötzinger complex may occur at a later stage. It is possible that ICV injections of STZ into the lateral ventricles may initially affect nuclei with proximity to the ventricle system. Similar to the position of the hippocampus relative to the lateral ventricles, the nTS is closely situated to the 4th ventricle and may thus be among the first brain areas exposed to STZ. However, it has been shown that brain regions further removed from the ventricles show various early (within 2–4 weeks) alterations that are induced by STZ and consistent with AD pathology [20 , 56]. These alterations include decreased antioxidant capacity (cortex, general brainstem tissue, cerebellum), as well as decreased energy metabolism, increased oxidative damage, and astrogliosis (cortex). In humans, deteriorations in brainstem areas (including the nTS) seem to be among the earliest events in the course of AD [10 , 57]. Furthermore, independent from a possible direct impact of STZ, other nuclei will be affected due to the interconnecting neuronal pathways. For example, the nTS is reciprocally connected to many brain regions [58 –62], including those directly impacted by AD [63 –67]. Thus, any perturbations in the nTS and its output will affect other brain centers.

AD patients experience respiratory deficits that manifest in sleep-disordered breathing [5, 8], blunted minute ventilation, and shortness of breath [9]. For the first time, we demonstrated respiratory alterations with ICV-STZ, a model that is commonly used to mimic AD of the sporadic type (95% of AD cases). The STZ-induced model seemed suited for this study as it closely mimics the progression of multiple symptoms and pathological hallmarks known from AD patients [14–16 , 68]. Here, we showed that ICV-STZ also induced markedly blunted responses to hypoxia by the peripheral chemoreflex and that minute ventilation was increased at rest. In a tauopathy model (Tau.P301L mice) that represents an important aspect of human AD, reduction of respiratory frequency and enlarged chest movements are observed in the terminal stage [69, 70]. These alterations are also evident during hypercapnia and associated with an abnormal serotonergic modulation of the respiratory network. A model of Parkinson’s disease (striatum injections of 6-OHDA), which is commonly associated with dementia of the AD-type [71], has significantly reduced respiratory frequency and minute ventilation [72]. Also, responses to hypercapnia (not hypoxia) are reduced and corroborated by neuronal loss in the ventral respiratory column. The changes documented in both models (Tau.P301L and 6-OHDA) cannot fully explain the respiratory abnormalities in STZ rats of this study. Although depressed respiration was observed in all three models, these changes were not apparent in the STZ model under baseline conditions. Furthermore, there were no changes in tidal volume and little effects on central chemoreflex responses to hypercapnia. It is still debated whether central chemoreception is an inherent property of many neurons within the dorsal and ventral respiratory column [73], or whether very few nuclei, foremost the retrotrapezoid nucleus (RTN), govern the central response to hypercapnia [74]. The impact of STZ-induced changes in the RTN (and/or other putative centers for central chemoreception) and on breathing parameters during hypercapnia may be limited two weeks following ICV injections.

The neuronal changes behind reduced peripheral chemoreflex function of STZ rats are not known. One possible mechanism may involve astrogliosis as observed in this study. In healthy conditions, reactive astrocytes form in response to stressors, such as inflammation and reactive oxygen species (ROS), and typically hold a protective role [34]. In the course of AD, however, it has been shown that astrogliosis contributes to inflammation and ROS formation due to an altered stress response (loss of a protective role) [33, 75]. Proinflammatory cytokines and ROS impair nTS function by inducing neuronal hyperexcitability [29, 76]. Additionally, astrogliosis in AD is associated with prolonged glutamatergic signaling due to an impaired clearance of glutamate in the synaptic cleft [33]. Hyperexcitability is observed in cortical brain region of AD patients [77, 78] and may thus also be present in the nTS via similar (yet unknown) mechanisms. Elevated output of respiratory centers could provide a basis for the increase in resting minute ventilation in STZ rats. Also, hyperexcitation of the nTS could limit effectiveness of sensory signals (ceiling effect) and lead to blunted chemoreflex function. In the long-run, persistent hyperexcitability may lead to excitotoxicity and cell death, similar to that observed in brains of AD patients [67]. However, whether increased excitation of nTS neurons accounts for the respiratory changes in STZ rats has to be addressed in future studies.

Analysis of sighs gives valuable information about the hypoxic state of the animal [40]. While sigh frequency is modulated by vagal afferents and chemoreceptors in the carotid body, sigh volume seems to be under the influence of peripheral chemoreceptors alone [79]. In STZ rats we measured a significantly augmented sigh volume in response to hypoxia. This change of sigh amplitude increases alveolar ventilation and may resemble efforts in the STZ rat to compensate for augmented hypoxemia due to a blunted chemoreflex. Alternatively, sigh amplitude may generally increase in the course of ICV-STZ. Such a change would be beneficial and opposite to the consequences of a blunted chemoreflex. However, with a constitutive alteration of sigh amplitude, a change in sigh amplitude would also be expected under baseline condition (21% O₂). Our data did not show changes of sigh amplitude under baseline in the STZ rat, which makes the alternative explanation less likely.

The pre-Bötzinger complex within the medullary respiratory network plays an important role in the control and generation of eupneic breaths and sighs [80]. Even though we did not find astrogliosis in the pre-Bötzinger complex, neuronal activity may still be altered with ICV-STZ. To date, it is unclear whether a distinct population of inspiratory neurons in the pre-Bötzinger complex forms the basis of sigh rhythmogenesis [81], or whether single neurons are able to simultaneously generate eupneic breathing and sighs [82]. In the latter, a single pattern-generating neuron produces rhythms for sighs and eupneic breaths, which is then modulated by the surrounding respiratory network. Both possibilities likely exist but the relative importance is unknown. Our data shows that two aspects of respiration are altered with ICV-STZ when challenged with hypoxia: breathing frequency was blunted (but not sigh frequency) and the volume of sighs (but not tidal volume of a regular breath) was augmented. Such specific modulation favors the importance of separate neurons responsible for the generation of sighs and regular breaths, since network modulation would change both parameters simultaneously when generated by a single neuron (e.g., cholinergic modulation: increase in sigh frequency and decrease of eupneic breath frequency) [82]. Importantly, this differential modulation also implies that STZ microinjections do not induce unspecific global changes, but rather differentially affect certain components of the respiratory system.

In summary, we demonstrated that ICV-STZ induces increased respiration at rest and blunted chemoreflex responses to hypoxia. These symptoms were corroborated by marked astrogliosis in the commissural nTS that were similar to those seen in the hippocampus. The results of this study also underline the versatility of this model utilizing ICV-injections of STZ, which has been extensively used in the past to elucidate the pathological hallmarks of memory and cognitive decline associated with clinical AD.

Footnotes

ACKNOWLEDGMENTS

We thank Dr. E.M. Hasser for allowing us to utilize the plethysmography chambers at the University of Missouri, and Drs. D.S. Middlemas and T. Walston (Truman State University) for the provision of the microscopes for our immunohistochemical analysis. We furthermore thank Drs. D.D. Kline and R.W. Baer for their valuable feedback on the manuscript.

Authors’ disclosures available online ().

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