The experimental protocols were approved by the Des Moines University Medicine and Health Sciences Institutional Animal Care and Use Committee and were conducted in accordance with the National Institutes of Health (NIH Publication No. 85-23, revised 1996) and the American Physiological Society’s Guide for the Care and Use of Laboratory Animals. The authors wish to declare that our work complies with the animal ethics checklist outlined by Grundy (Grundy, 2015).

Adult male Sprague-Dawley rats (Envigo, Madison, WI) of similar age (8–10 weeks) and weight (250–300 g) were used for these experiments. All rats were group-housed under controlled temperature (23°C) and humidity conditions and kept on a 12:12 h light-dark cycle and fed standard rat chow with water available ad libitum. At the beginning of the study 30 animals were randomly assigned to the following groups (n = 15 per group): Sham (animals exposed to air-air cycling), CIH (animals exposed to intermittent hypoxia-air cycling). In certain cases we were unable to use a given data stream or in other cases not all animals were used for all experiments. For each experimental variable, the n values are given in the results section as well as the figure legends. Both groups followed identical experimental timelines (shown in Figure 1).

Both experimental groups were exposed to 14 sequential days of chronic intermittent hypoxia (CIH, 60 s F_iO₂ 0.10, 120 s F_iO₂ 0.21) or normoxia/normoxia cycling (sham, 60 s F_iO₂ 0.21, 120 s F_iO₂ 0.21), as previously described (Marcus et al., 2009; Marcus et al., 2010; Philippi et al., 2010; Marcus et al., 2012). The CIH protocol ran for 8 h per day (from 07:30-15:30) and took place in a pair of adjacent Plexiglas chambers (Coy Laboratory Products Inc., Glass Lake, MI) in a designated room in the animal care facility. All animals remained in their home cages which were placed in the Plexiglas chambers during daily exposures.

At the conclusion of the CIH or sham protocol, GFR was assessed non-invasively in conscious, unrestrained rats by measuring clearance of the fluorescent tracer FITC-sinistrin as described previously (Schock-Kusch et al., 2009; Schock-Kusch et al., 2011; Friedemann et al., 2016; Herrera Pérez et al., 2016; Kious et al., 2022). Briefly, FITC-sinistrin clearance was measured using the preclinical transdermal GFR monitor (Medibeacon, St. Louis, MO). On the day prior to the study, a site on the back was shaved and prepared with a depilatory agent (i.e. Nair). On the day of the study the rats were briefly anesthetized (1.5% Isoflurane in air) and the depilated area was cleaned. The monitor was affixed to the back with an adhesive patch and a bolus injection of FITC-sinistrin (5 mg/100 g BW) (Herrera Pérez et al., 2016) was given through one of the lateral tail veins. Measurements of FITC-sinistrin fluorescence were made once per second for 120 min. At the conclusion of the experiment, the monitor and battery pack were removed, and the data was downloaded for later analysis with MPD Lab software (Medibeacon, St Louis, MO) as described previously (Friedemann et al., 2016).

Experiments to measure RBF, RVC, RP, CPO₂, and MPO₂ were conducted 24 h after GFR measurements. For surgical physiological experiments, anesthesia was induced with 4% inhalation isoflurane in air after which time anesthesia was reduced to 1.5%–2.0% as necessary to maintain a surgical plane of anesthesia. In order to determine appropriate anesthetic depth was reached, a toe pinch and/or tail pinch was administered after induction and every 15 min thereafter. The absence of paw withdrawal, movement of the tail, vocalization, or marked increase in respirations after a pinch was taken as evidence of adequate anesthetic depth. Body temperature was measured using a rectal thermometer and monitored using a Physiosuite w/RightTemp Temperature Monitoring and Homeothermic Control system (Kent Scientific, Torrington, CT), and was maintained at 37°C using a far infrared heating pad integrated with a Surgisuite surgical platform (Kent Scientific, Torrington, CT).

After appropriate anesthetic depth was reached, the left femoral artery was cannulated with a Millar Mikro-Cath pressure catheter (Millar, Houston, TX) connected to a bridge amp (AD Instruments, Colorado Springs, CO) for measurement of arterial pressure. Then a ventral mid-line incision was made, and the kidney was approached in the peritoneal space. Once the renal artery and vein were identified the renal artery was gently separated from the vein and a transit-time flow probe (Transonic, Ithaca, NY) was placed around the renal artery. After the flow probe was secured, two needle probes were inserted 0.5–1 mm (cortical) and 3–4 mm (medullary) into the kidney to measure tissue PO₂ using the Oxylite Pro (Oxford Optronix, United Kingdom). In a sub-set of CIH and sham animals (n = 7 per group), a laser speckle contrast imager (Moor FLPI-2, Moor Instruments, Devon United Kingdom) was positioned with the whole kidney in the field of view for assessment of renal microcirculatory perfusion (RP) (in addition to RBF, CPO₂, and MPO₂). After a 60-min equilibration period, a 5-min baseline period of RBF, arterial pressure, RP, and cortical and medullary PO₂ was recorded for analysis. After measurement of baseline values, a series of ten 30 s episodes of intermittent asphyxia (IA) were initiated. Each IA episode was characterized by an F_iO₂ of 0.10 and F_iCO₂ 0.03 with the remaining balance made up of nitrogen. Each IA episode was followed by a 60 s recovery period breathing medical grade air (F_iO₂ 0.21, balance nitrogen). All variables were measured continuously during the IA protocol.

At the conclusion of the experimental protocol all rats were humanely euthanized. In accordance with standards set forth by the American Veterinary Medical Association and the Institutional Animal Care and Use Committee at Des Moines University Medicine and Health Sciences, all rats were euthanized via anesthetic overdose (5% isoflurane) combined with the addition of a secondary physical method of euthanasia.

At the conclusion of physiological experiments and after anesthetic overdose, rat kidneys were excised and dissected, flash frozen in liquid nitrogen, and stored at −80°C for later analysis. Studies of renal mRNA expression were performed as previously described (Kious et al., 2022). Briefly, RNA isolation and cDNA synthesis were performed according to manufacturer instructions using the Direct-zol RNA MiniPrep Plus (Zymo, Irvine, CA) and High-Capacity cDNA Reverse Transcriptase (Thermo Fisher Scientific, Waltham, MA) kits, respectively. Gene expression was assessed by Sso Advanced SYBR green chemistry (Bio-Rad, Hercules, CA) real-time PCR following reverse transcription of total RNA. Real-time PCR was performed on a CFX Connect Real Time PCR Detection System (Bio-Rad, Hercules, CA). β-Actin mRNA was quantified as an internal control for each sample and quantifications were performed using the ΔΔCt method and expressed as fold change. Forward and Reverse primer sequences are show in Table 1. The following genes were probed in our samples: Krüppel-like factor 2 (KLF2), nuclear factor erythroid 2-related factor 2 (NRF2), copper zinc superoxide dismutase (CuZn SOD), NAD(P) H oxidase catalytic subunit (gp91phox), Krüppel-like Factor 15 (KLF15), E Cadherin, connective tissue growth factor (CTGF), tissue inhibitor of metalloproteinase 1 (TIMP), vascular endothelial growth factor (VEGF), Galectin-3, snail family zinc finger 1 (snail), and collagen I.

All physiological parameters except GFR (MPD Lab, Medibeacon, St Louis, MO) and RP (moorFLPI2 Research Software, Moor Instruments, Devon, United Kingdom) were recorded with a Powerlab data acquisition and analysis system (AD Instruments, Colorado Springs, CO). Mean values for baseline (B1) RBF, RP, and CPO₂ and MPO₂ were calculated from 5-min of data immediately following a 60-min post-instrumentation equilibration period (pre-IA baseline). In addition to computing baseline values, we quantified the RBF, RP, and CPO₂/MPO₂ response to IA using the following approaches. Baseline (b2-b10) and nadir (n1-n10) measurements occurring during IA were based on 10 s periods centered around appropriate peaks and nadirs. First, we analyzed the change (if any) in normoxic baseline RBF, RP, CPO₂, and MPO₂ relative to the pre-IA baseline (e.g. B1-B2, B1-B3, B1-B4, etc.). Then we analyzed the average change in RBF, RP, and CPO₂/MPO₂ during the IA protocol (e.g. B1-N1, B2-N2, B3-N3, etc.). A graphical representation of the time points analyzed is shown in Figure 2. Group mean data was analyzed using unpaired t-tests. All data was analyzed using GraphPad Prism version 9.5.1 for Windows software (San Diego, CA), and the threshold for statistical significance was set at p < 0.05. Values in the results section are expressed as mean ± standard deviation, and in the figures as median and quartiles with violin plots.

As shown in Figure 3, after 2 weeks of CIH or sham conditioning, glomerular filtration rate (GFR) in conscious unrestrained rats was significantly higher in the CIH group than the sham group (1.09 ± 0.23 ml/min/g BW sham vs. 1.39 ± 0.34 ml/min/g BW CIH, t 2.29 df 18, n = 10 sham and CIH, p = 0.03). Baseline hemodynamic, CPO₂, and MPO₂ values are shown in Table 2. RBF (8.2 ± 3.3 ml/min sham vs. 4.6 ± 2.1 ml/min CIH, t 3.45 df 27, n = 15 sham, n = 14 CIH, p = 0.002), RVC (0.12 ± 0.03 ml/min/mmHg sham vs. 0.07 ± 0.05 ml/min/mmHg CIH, t 2.81 df 23, n = 13 sham, n = 12 CIH, p = 0.005), CPO₂ (41.5 ± 8.4 mmHg sham vs. 31.0 ± 8.2 mmHg CIH, t 3.14 df 23, n = 14 sham, n = 11 CIH, p = 0.005), and MPO₂ (29.8 ± 7.5 mmHg sham vs. 18.1 ± 2.5 mmHg CIH, t 5.71 df 25, n = 12 sham, n = 15 CIH, p = 0.005) were all lower in CIH vs sham. In contrast, arterial pressure was significantly higher in CIH vs sham animals (73.2 ± 14.3 mmHg sham vs. 87.1 ± 14.9 mmHg CIH, t 2.45 df 25, n = 15 sham, n = 12 CIH, p = 0.021). Heart rate was not significantly different between groups at baseline (315.3.2 ± 5.9 mmHg sham vs. 307.0 ± 35.6 mmHg CIH, t 0.74 df 25, n = 15 sham, n = 12 CIH, p = 0.021).

Hemodynamic and tissue PO₂ responses to the IA protocol are shown in Figures 4–6. Normoxic RBF (-0.43 ± 0.54 ml/min sham vs. -0.98 ± 0.59 ml/min CIH, t 2.2 df 18, n = 10 per group, p = 0.04), RVC (-0.0006 ± 0.008 ml/min/mmHg sham vs. -0.006 ± 0.006 ml/min/mmHg CIH, t 1.6 df 18, n = 10 per group, p = 0.06), RP (-0.84% ± 3.6% sham vs. -4.33 ± 1.9% CIH, t 2.26 df 12, n = 7 per group, p = 0.04) CPO₂ (-2.4 ± 1.4 mmHg sham vs. -4.1 ± 2.2 mmHg CIH, t 2.2 df 20, n = 11 per group, p = 0.04) and MPO₂ (-2.8 ± 3.7 mmHg sham vs. -6.5 ± 3.6 mmHg CIH, t 2.2 df 18, n = 10 per group, p = 0.04) baselines during IA decreased relative to pre-IA baseline in both groups, but to a greater extent in CIH. Similarly, RBF (-0.41 ± 0.22 ml/min sham vs. -0.75 ± 0.45 ml/min CIH, t 2.2 df 13, n = 10 per group, p = 0.04), RVC (-0.002890 ± 0.006101 ml/min/mmHg sham vs. -0.01488 ± 0.01473 ml/min/mmHg CIH, t 2.4 df 18, n = 10 per group, p = 0.03), RP (-4.6 ± 1.3% sham vs. -8.4 ± 1.3% CIH, t 5.7 df12, n = 7 per group, p = 0.0001) and CPO₂ (-9.0 ± 3.2 mmHg sham vs. -13.3 ± 5.0 mmHg CIH, t 2.4 df 20, n = 11 per group, p = 0.02) MPO₂ (-5.8 ± 3.2 mmHg sham vs. -10.7 ± 5.0 mmHg CIH, t 2.6 df 18, n = 10 per group, p < 0.02) decreased during asphyxia in both groups, but to a greater extent in CIH vs sham.

Results of qRT-PCR studies on renal cortical tissue are shown in Figure 7. Expression of genes related to antioxidant defense (KLF2 (2.81 ± 1.9-fold change sham vs. 1.46 ± 0.89-fold change CIH, t 2.2 df 19, n = 9–12 per group, p = 0.04) NRF2 (0.59 ± 0.26-fold change sham vs. 0.28 ± 0.19-fold change CIH, t 2.5 df 12, n = 6–8 per group, p = 0.03), CuZnSOD (0.93 ± 0.33-fold change sham vs. 0.55 ± 0.30-fold change CIH, t 2.2 df 13, n = 5–10 per group, p = 0.04) were downregulated while genes associated with the pro-oxidative catalytic subunit of NADPH oxidase (gp91phox (1.44 ± 1.8-fold change sham vs. 4.54 ± 4.2-fold change CIH, t 1.9 df 14, n = 8 per group, p = 0.04) were upregulated in CIH vs. sham. Expression of anti-fibrotic genes KLF15 (1.10 ± 0.35-fold change sham vs. 0.62 ± 0.44-fold change CIH, t 2.4 df 15, n = 8–9 per group, p = 0.02), and E-Cadherin (1.03 ± 0.54-fold change sham vs. 0.32 ± 0.25-fold change CIH, t 2.8 df 9, n = 5–6 per group, p = 0.02) were lower in CIH vs sham, whereas expression of pro-fibrotic/fibrosis-associated genes (CTGF (1.29 ± 0.51-fold change sham vs. 2.10 ± 0.79-fold change CIH, t 2.3 df 13, n = 7–8 per group, p = 0.04), TIMP (1.13 ± 0.22-fold change sham vs. 2.04 ± 0.83-fold change CIH, t 2.4 df 14, n = 7–9 per group, p = 0.03), VEGF (1.22 ± 1.13-fold change sham vs. 2.35 ± 1.62-fold change CIH, t 1.8 df 18, n = 9–11 per group, p = 0.04), Galectin 3 (0.81 ± 0.54-fold change sham vs. 2.08 ± 1.74-fold change CIH, t 1.9 df 12, n = 7 per group, p = 0.04), snail (0.91 ± 0.37-fold change sham vs. 2.02 ± 1.38-fold change CIH, t 2.2 df 14, n = 8 per group, p = 0.04), and collagen I (0.91 ± 0.60-fold change sham vs. 1.86 ± 1.05-fold change CIH, t 2.2 df 15, n = 7–10 per group, p = 0.04) were higher in CIH vs. sham.

We observed that GFR was higher in CIH relative to sham, an observation consistent with the hyperfiltration observed in patient populations with SA (Kinebuchi et al., 2004; Adeseun and Rosas, 2010). In contrast, O’Neil et al. found a lower GFR in CIH conditioned rats relative to sham (O'Neill et al., 2019), an observation which is also consistent with clinical studies showing reduced GFR in patients with SA (Li et al., 2019). Given the similarities of the intensity and duration of the CIH paradigms used in our study and that of O’Neil et al. it is difficult to reconcile these differences. It may be that our findings represent a hyperfiltration state which precedes eventual decreases in GFR associated with prolonged or more severe CIH exposure. Progression from hyperfiltration to reduced GFR occurs in diabetic nephropathy (Palatini, 2012), so this phenomenon is not without precedent.

Another explanation may be the physiological state under which GFR was measured in these two studies. In our study, GFR was measured in conscious unrestrained animals, whereas the study by O’Neil et al. measured GFR in pentobarbital anesthetized rats (O'Neill et al., 2019). Previous studies have shown that pentobarbital anesthesia in rats causes a marked reduction in mean arterial pressure, heart rate, GFR, and effective renal plasma flow (Walker et al., 1986). Determining whether or not the differences in the effect of CIH on GFR observed between studies can be attributed to anesthesia will require future studies to perform both techniques in the same animals. While our studies do not directly address mechanisms of glomerular hyperfiltration after CIH conditioning there are multiple physiological changes occurring with CIH that may contribute. Vascular and autonomic dysfunction associated with CIH are mediated in part by chronic RAAS activation (Marcus et al., 2010; Marcus et al., 2012), which has been shown to elicit constriction of the efferent arterioles and increase GFR (Cortinovis et al., 2022). Further study is needed to fully address the nature and time course of changes in GFR with CIH conditioning.

The present study shows significantly lower baseline RBF, RVC, and RP as well as exaggerated hypoxia-evoked reductions in RBF, RVC, and RP in CIH relative to sham. The only study that has previously explored the effect of CIH conditioning on baseline RBF found no significant difference in RBF or renal vascular resistance (RVR) between CIH and sham groups (O'Neill et al., 2019). This unexpected finding may be explained by the nature of CIH related changes in control of organ blood flow. CIH contributes to enhanced baseline sympathetic nerve activity and increased arterial pressure (Huang et al., 2009; Gilmartin et al., 2010; Marcus et al., 2010; Del Rio et al., 2016), as well as enhanced sympathetic responses to hypoxic events (Huang et al., 2009; Gilmartin et al., 2010; Marcus et al., 2010). CIH specifically augments chemoreflex control of renal sympathetic nerve activity (Huang et al., 2009) (RSNA), and previous work has established that conditions characterized by chemoreflex-mediated increases in RSNA have the potential to cause reductions in RBF and RVC, and increases in RVR (Clayton et al., 2011; Marcus et al., 2015; Pügge et al., 2016; Kious et al., 2022). Thus in a CIH model it would be logical to hypothesize reductions in RBF and RVC. Both our study and the study by O’Neil et al. measured RBF under anesthesia, however as mentioned previously, O’Neil et al. used pentobarbital anesthesia which has been shown to obtund cardiovascular reflex function (Peiss and Manning, 1964). Therefore, it is possible that pentobarbital suppressed chemoreflex control of RSNA and for this reason the expected reductions in RBF after CIH were not observed by O’Neil et al. In addition to changes in autonomic function that affect control of blood flow, vascular dysfunction also likely plays a role in the dysregulation of integrated hemodynamic control we observed. We have previously shown that CIH is associated with vascular endothelial dysfunction (Philippi et al., 2010; Marcus et al., 2012) and oxidative stress (Marcus et al., 2010; Philippi et al., 2010), however these parameters have not previously been assessed in the renal vasculature in the CIH model. ROS may be generated in CIH animals as a results of mitochondrial dysfunction (Prabhakar et al., 2007), or acutely in response to hypoxia, angiotensin II (ANGII), and/or proinflammatory cytokines (Liu et al., 2017). CIH is associated with RAAS activation, renal inflammation, and oxidative damage in the kidney (Sun et al., 2012). ROS in the kidney has previously been shown to reduce renal medullary blood flow and to induce afferent arteriolar constriction in presence of ANGII (Ratliff et al., 2016). Thus, the multiple pathophysiological insults associated with CIH are consistent with established causes of autonomic and vascular dysfunction, and the observed dysregulation of renal hemodynamic control observed in our studies.

Previous studies have shown a reduction in resting normoxic renal cortical but not medullary tissue PO₂ after conditioning with a similar CIH paradigm (O’Neill et al., 2019). Interestingly, this occurred in the presence of a reduction in GFR and sodium transport without a reduction in renal artery blood flow. In addition, the authors noted that CIH animals exhibited higher renal oxygen consumption at any given level of sodium transport. Thus, in this study decreases in cortical tissue PO₂ were driven by changes in tissue metabolism rather than as a result of decreased oxygen delivery. In the current study, both cortical and medullary PO₂ were lower in CIH exposed animals at baseline and during the IA protocol. While the reason for this discrepancy is unclear, one potential explanation might center around differences in GFR between the studies. In our study we observed increased GFR and decreased RBF in CIH animals under normoxic conditions, whereas O’Neil et al. observed decreased GFR and sodium transport in CIH animals relative to sham. While O’Neil et al. observed no differences in oxygen consumption between CIH and sham, this was in the face of lower GFR and sodium transport. It is possible that a higher GFR in our study drove greater sodium transport and oxygen consumption contributing to lower tissue PO₂. We did not measure tissue oxygen consumption or sodium transport, so our data can neither directly support nor refute this possibility, but it is a plausible explanation based on the data we collected.

Under normal circumstances, renal tissue PO₂ is maintained at stable levels by a unique interaction between RBF, GFR, and oxygen consumption (QO₂) (Brezis et al., 1994). Renal QO₂ is primarily driven by the metabolic demands associated with tubular solute reabsorption, however this relationship can become dysregulated as a result of mitochondrial dysfunction, RAAS activation, and hypoxic conditions (Brezis et al., 1994; Haase, 2013). Disease states such as diabetes and hypertension are often associated with increased QO₂ and decreased renal tissue PO₂ (Hansell et al., 2013). In these diseases, elevated kidney QO₂ is commonly mediated by increased angiotensin receptor tonus and oxidative stress which contributes to progression of renal damage and reduced function (Hansell et al., 2013). Given the well-known RAAS activation and oxidative stress in CIH models of SA, these factors likely contribute to the decreases we observed in renal tissue PO₂. Thus, the combination of oscillations in RSNA and RBF/RP combined with persistent RAAS activation and intermittent hypoxemia (associated with individual episodes of asphyxia) act synergistically to create a state of tissue hypoxia.

Because we observed notable decreases in RBF/RVC/RP in CIH conditioned animals, we sought to determine how this might affect gene expression of transcription factors related to blood flow. We measured expression of the shear stress-sensitive transcription factor Krüppel-like factor 2 (KLF2), as KLF2 controls expression of mediators of antioxidant defense, nitric oxide production, and tissue fibrosis (Dekker et al., 2005; Boon et al., 2007; Fledderus et al., 2008) and has previously been shown to be downregulated in animal models characterized by reduced RBF (Haack et al., 2014; Del Rio et al., 2017; Marcus et al., 2018; Kious et al., 2022). As shown in Figure 7, we observed that the gene expression of renal cortical KLF2 and the master antioxidant transcription factor NRF2 (a downstream target of KLF2) were significantly downregulated in CIH relative to sham. To determine if NRF2 downregulation resulted in decreased gene expression of antioxidant enzymes under its control, we measured mRNA expression of CuZn SOD, which was also significantly decreased in CIH vs sham. In contrast, gene expression of the catalytic subunit of NADPH oxidase (gp91phox) was significantly increased in CIH vs sham, consistent with findings in other studies (Prabhakar et al., 2007; Sun et al., 2012).

Tissue fibrosis has been theorized to be a common pathway to renal dysfunction (Wang et al., 2022) and may resist oxygen diffusion and contribute to tissue hypoxia (Edwards and Kurtcuoglu, 2022), so we examined gene expression of fibrosis related pathways in renal cortical tissue. In particular we focused on pathways related to tissue hypoxia because of its relevance to our physiological measurements. KLF15, and E-Cadherin, which have fibrosis-suppressing activity and have been shown to be downregulated by ANG II and hypoxia (Esteban et al., 2006; Lu et al., 2019), were decreased in cortical tissue from CIH relative to sham. Expression of pro-fibrotic CTGF, TIMP, Galectin 3, Snail, and VEGF (all of which are induced by hypoxia and/or ANG II (Liu et al., 2017; Boutin et al., 2022; Khalaji et al., 2023)), was significantly higher in cortical tissue from CIH vs sham, suggesting that CIH exposure promotes activation of pro-oxidative, pro-fibrotic signaling that may contribute to tissue damage and organ dysfunction.

Often, discerning the effects of multiple-concurrent co-morbidities or conditions can make clear interpretation of data difficult. The common presence of obesity, metabolic syndrome, and advanced age alongside sleep apnea, can confound interpretation of the relative contribution of any given variable. Our animal model examines the physiological changes that occur in response to CIH conditioning in the absence of common co-morbidities, giving a clearer picture of the contribution of CIH per se. These findings yield significant insight into the potential for the CIH associated with sleep apnea to precipitate or hasten the decline of renal function and to identify the mechanisms by which this might occur. In addition to the specificity of our model, our approach also has significant advantages. In this study we combine measurement of renal artery blood flow using a transit time flow probe with measurement of renal microcirculatory perfusion using laser speckle contrast imaging (LSCI). The larger imaging field used in LSCI allows for a more comprehensive quantification of ‘whole organ’ regulation of microcirculatory perfusion.

In this study direct measurement of RBF, RVC, RP, CPO₂, and MPO₂ is made at the conclusion of the experimental period under anesthesia. The use of anesthesia during invasive experiments is a limitation, however, any changes induced by anesthesia would be expected to affect both experimental and control animals, thus we do not feel this confounds interpretation of differences observed between groups. Measurement of physiological variables under anesthesia has inherent limitations regardless of anesthetic, but we feel that the difference in anesthetics used in similar studies (O’Neill et al., 2019) should be considered when comparing data. In addition, we acknowledge that the length of our CIH exposure (2 weeks) is short relative to the amount of time that patients with sleep apnea are typically exposed to CIH associated with their condition. Previous studies in rats and mice have shown that short-term CIH conditioning is sufficient to elicit enhanced chemoreflex control of sympathetic nerve activity (Huang et al., 2009; Marcus et al., 2010), however longer-term studies might reveal more profound effects of CIH on renal perfusion, tissue PO₂, and injury. Indeed, studies focused on dose response of CIH exposure time show marked changes in tissue redox regulation and fibrosis with longer exposure times (Sun et al., 2012).

Finally, in the studies presented we only assessed GFR, renal hemodynamic regulation, and tissue PO₂ in male animals, therefore inferences based on our study should be limited to the male sex. It is unclear how female sex steroid hormones interact with the insults associated with CIH, and therefore the effect of CIH on renal function and hemodynamic control in females deserves study.