No Benefit of 3% Hypertonic Saline Following Experimental Intracerebral Hemorrhage

Abstract

Intracerebral hemorrhage (ICH) is a stroke subtype with a high mortality rate (~ 40%). After ICH, the mass effect of the hematoma and edema contribute to raised intracranial pressure (ICP) and poor outcome. Endogenous compensatory mechanisms that blunt ICP elevations include redirection of venous blood and cerebrospinal fluid, along with brain tissue compliance (e.g., decreased cell volume, increased cell density); however, these limited reserves can be exhausted after severe stroke, resulting in decompensated ICP that requires careful clinical management. Management strategies can include administration of hypertonic saline (HTS), an osmotic agent that putatively attenuates edema, and thereby ICP elevations. Evidence regarding the efficacy of HTS treatment following ICH remains limited. In this study, adult male rats were given a collagenase-induced striatal ICH and a bolus of either 3% HTS or 0.9% saline vehicle at 2- and 14-hours post-stroke onset. Neurological deficits, edema, ipsilateral cell volume and density (in areas S1 and CA1), and contralateral CA1 ultrastructural morphology were assessed 24 h post-ICH. Animals had large bleeds (median 108.2 µL), extensive edema (median 83.9% brain water content in ipsilateral striatum), and evident behavioural deficits (median 5.4 neurological deficit scale score). However, HTS did not affect edema (p ≥ 0.4797), behaviour (p = 0.6479), cell volume (p ≥ 0.1079), or cell density (p ≥ 0.0983). Qualitative ultrastructural assessment of contralateral area CA1 suggested that HTS administration was associated with paradoxical cellular swelling in ICH animals. Overall, there was no benefit with administering 3% HTS after ICH.

Supplementary Information

The online version contains supplementary material available at 10.1007/s12640-024-00722-2.

Introduction

Intracerebral hemorrhage (ICH), where a blood vessel ruptures within the brain, accounts for ~ 10–15% of all strokes and has a devastating mortality rate of ~ 40% (Shoamanesh et al. 2021; Joundi et al. 2021). Currently, no clinically approved neuroprotective treatment for ICH exists, necessitating further research in the area. Following the primary injury exerted by the hemorrhage itself, secondary damage mechanisms continue to evolve in the days to weeks following stroke. One such mechanism is edema. With severe ICH, the added intracranial mass of the hematoma and associated edema may raise intracranial pressure (ICP), which independently exacerbates secondary injury and may further aggravate edema (Wilkinson et al. 2020). While there has been much clinical focus on elucidating the link between edema, ICP, and outcome (Marchina et al. 2023), comparably less research is focused on investigating the treatments used to manage edema and ICP (Chugh et al. 2021).

Multiple forms of edema occur after ICH, which are explored at length in other reviews (Ironside et al. 2019; Chen et al. 2021; Jiang et al. 2022). Most relevant to this study are cytotoxic and ionic edema, both of which begin within minutes to hours after ICH (Chen et al. 2021). Following ICH, mechanical injury and metabolic dysfunction in perihematomal regions impairs primary ion transport, facilitating the formation of cytotoxic edema (i.e., cellular swelling) as concentration gradients are lost. When left unchecked, cytotoxic edema compromises cellular health (Simard et al. 2007; Chen et al. 2021; Jiang et al. 2022). The movement of water into the intracellular space depletes extracellular water, creating an osmotic gradient that drives an influx of water from the intravascular space into the interstitial space; this is called ionic edema, which drives the initial swelling of brain tissue following stroke (Simard et al. 2007; Jiang et al. 2022). As hematoma resolution and the neuroinflammatory response progress, other forms of edema develop that worsen brain swelling, including serum extrusion and vasogenic edema (Ironside et al. 2019; Chen et al. 2021). These electrolyte and water imbalances directly impede brain function in perihematomal regions, and their impact likely extends farther, owing to the consequences of elevated ICP (Kalisvaart et al. 2020).

Several compensatory mechanisms are engaged in response to rising ICP in settings of acute mass effect; classically, these included the redirection of venous blood and cerebrospinal fluid (CSF) out of the brain, among others, while brain tissue itself was thought to be non-compressible (Wilkinson et al. 2020). However, our lab has shown that brain “tissue compliance” occurs after moderate to severe strokes in rodents: cells in regions distal to the bleed transiently decrease in volume and increase in packing density over the acute post-stroke period, presumably to compensate for pathologically elevated ICP (Kalisvaart et al. 2020; Wilkinson et al. 2023). When the intracranial volume reserves afforded by these compliance mechanisms are exhausted or impeded (which often occurs following severe stroke (Wilkinson et al. 2020), ICP rises in a decompensated fashion, requiring careful clinical management to mitigate risk of additional brain injury or patient mortality due to brainstem compression (Shoamanesh et al. 2021). While routine management of ICP is not recommended after ICH (Shoamanesh et al. 2021), those showing signs of decompensated ICP (e.g., vomiting, papilledema, pupillary dilation) must be carefully monitored, as persistently elevated ICP is associated with increased risk of mortality, partly due to brainstem herniation, and worse functional outcome (Chen et al. 2019; Wilkinson et al. 2020). Clinical management of ICP in such circumstances range from simple interventions, such as head elevation, to more invasive measures, such as decompressive craniectomy (Shoamanesh et al. 2021).

Given that edema contributes to mass effect following stroke, one might assume that the magnitude of edema should relate to acute ICP dynamics. Therefore, edema-reducing treatments are occasionally used to manage elevated ICP; indeed, osmotic agents like hypertonic saline (HTS) are tentatively recommended (Level of Evidence C) as a temporizing measure for elevated ICP following ICH (Shoamanesh et al. 2021). Though HTS is believed to work through multiple mechanisms of action, it is primarily thought to reduce edema via osmotic mechanisms (Forsyth et al. 2008; Fink 2012; Barik et al. 2023). By increasing the osmolarity of blood, HTS counteracts cytotoxic and ionic edema; in doing so, it presumably attenuates elevated ICP by drawing excess fluid out of the brain. While several animal studies have found benefit with HTS administration, there remains little direct evidence supporting the administration of HTS after ICH in human patients (Kamel et al. 2011; Leasure et al. 2016; Shah et al. 2018; Shoamanesh et al. 2021; Holden et al. 2023). This is perhaps due to a lack of consensus around the ideal concentration, dose, timing, and even method of administration of HTS (Kamel et al. 2011; Fink 2012; Barik et al. 2023; Holden et al. 2023). Alternatively, there are a few additional explanations for such ambiguity. First, HTS has not consistently reduced edema or improved functional outcome in clinical settings (Qureshi et al. 1999; Schwarz et al. 2002; Brogan and Manno 2015; Shah et al. 2018); a recent systematic review found that while HTS was associated with improved ICP control in clinical studies of ischemic stroke, this was not accompanied by improvements in edema (Chugh et al. 2021). Second, there is limited evidence that edema is an independent predictor of outcome post-ICH (Wilkinson et al. 2020; Kalisvaart et al. 2023; Marchina et al. 2023); a recent clinical meta-analysis found only a weak relationship between edema and outcome after ICH (Marchina et al. 2023). Lastly, it is unclear whether reductions in edema truly lead to meaningful reductions in ICP (Wilkinson et al. 2020; Kalisvaart et al. 2023; Marchina et al. 2023); this lack of a clear linear relationship between edema and ICP may be explained in part by the contribution of ICP compliance mechanisms, such as tissue compliance (Wilkinson et al. 2020). Therefore, HTS administration may not only affect edema, but also tissue volume throughout the brain.

Given that ionic and osmotic homeostasis are intricately tied to cell volume regulation, administration of HTS may augment tissue compliance following stroke. By establishing an osmotic gradient in which water is drawn from the interstitial to intravascular compartments, cell volume may be further reduced via activation of osmo-sensitive and mechanosensitive ion channels, accentuating the cellular efflux of water and ions (e.g., K⁺, Cl⁻) into the interstitial compartment (Hoffmann et al. 2009). However, enhancing tissue compliance may come with risk; because cell volume and function are intimately tied (Danziger and Zeidel 2015), HTS may ultimately have a detrimental effect on cell health, which has not been previously studied after ICH. Our preceding work indicates that tissue compliance within uninjured brain regions is associated with subcellular signs of stress, such as edematous changes to organelles and evidence of mitochondrial lysosomal transition (Kalisvaart et al. 2020). Additionally, other risks of HTS administration include hyperchloremic acidosis, rebound ICP elevations, and in instances of overly rapid correction, osmotic demyelination syndrome (ODS) (Fink 2012; Bouchat et al. 2019). Considering these risks and the lack of certainty regarding the effect of HTS on edema and stroke outcomes, further study of this issue is crucial. While HTS remains a candidate for attenuating edema, reducing ICP, and improving outcome, more in-depth research is needed to clarify the widespread effects this treatment could have.

To investigate these hypotheses, we compared bolus dosing of 3% HTS to 0.9% saline (SAL) vehicle in rats, beginning 2-hours following collagenase-induced striatal ICH, and then 12-hourly until euthanasia. While previous preclinical ICH studies have used higher concentrations, 3% HTS is commonly used in clinic (Zeynalov et al. 2008; Wagner et al. 2011; Shoamanesh et al. 2021; Jiang et al. 2022; Holden et al. 2023). The collagenase model is widely used, and it reproduces mechanisms that drive edema formation in humans (MacLellan et al. 2008; Manaenko et al. 2011; Kalisvaart et al. 2023). In Exp. 1, we hypothesized that HTS administration would not affect hematoma volume at 8 h post-ICH, which is an important safety check. In Exp. 2, we hypothesized that HTS would reduce edema compared to SAL-treated rats. In Exp. 3, we hypothesized that HTS would augment tissue compliance (decreased cell volume and increased cell packing density compared to saline-treated controls) and worsen cellular health assessed via transmission electron microscopy (TEM), both at 24 h post-ICH. The 24-hour timepoint was chosen as it is a time of significant cytotoxic and ionic edema (Ironside et al. 2019; Chen et al. 2021). We additionally predicted that HTS administration would improve behavioural outcomes.

Materials and Methods

Ethics

All experiments were conducted according to the Canadian Council on Animal Care guidelines and were approved by the University of Alberta’s Biosciences Animal Care and Use Committee (protocol: AUP 960). All animals were closely monitored following surgery until euthanasia.

Subjects

One hundred male Sprague Dawley rats (~ 3–4 months old for all experiments; 250–350 g for Exps. 1 and 3; 300–400 g for Exp. 2) from Charles River Laboratory (Saint Constant, Quebec) were used. Rats were housed 3 to 4 per cage pre-surgery and individually housed post-ICH surgery, in a temperature-controlled environment on a 12-hour light cycle. Food (Purina rodent chow) and water were given ad libitum. All animals were kept in the same location and were not housed according to treatment group.

Experimental Design

The study was conducted as specified in an a priori planning document (Online Resource 1). Statistical power was calculated to allow for 80% power in our primary endpoint across all experiments, and statistical tests were pre-planned to minimize bias (see Statistical Analyses section below). Any deviations from our plan are noted in the text. Sterile solutions of 3% HTS or normotonic 0.9% saline (SAL) were administered to the lateral teil vein while under isoflurane anesthetic. For Exp. 1, a single dose of HTS or SAL was administered at 2-hours post-ICH, and for Exps. 2 and 3, two doses of HTS or SAL were administered, one each at 2- and 14-hours post-ICH. A graphical depiction of experimental timelines is provided (Fig. 1). All animals were assigned to groups using a random number generator (random.org). For all experiments, researchers were blinded by a third experimenter for both treatment (HTS vs. SAL) and, where applicable, surgery (ICH vs. SHAM). Specifically, unlabelled syringes of SAL or HTS were prepared by one researcher, and administered by another who was blinded to group identity for all experiments. For Exp. 3, coded vials of collagenase or 0.9% saline (ICH vs. SHAM) were prepared by a separate researcher to ensure the surgeon was blinded to group identity.

Fig. 1.

Timeline of Experiments Experimental timelines for all experiments are shown. Shaded bars indicate bolus dosing of HTS (3% hypertonic saline) or SAL (0.9% saline). Hematoma volume, via hemoglobin assays, and pain, via the Rat Grimace Scale, were assessed in Exp. (1). Global edema, via wet-weight dry-weight, and behaviour, via the Neurological Deficit Scale, were assessed in Exp. (2). Cell volume and cell packing density, along with ultrastructural morphology and NDS were assessed in Exp. (3). HemV = Hematoma Volume, RGS = Rat Grimace Scale, NDS = Neurological Deficit Scale, CV = Cell Volume, CD = Cell Density, Ultrastructure = Ultrastructural morphology. ICH animals received a striatal injection of collagenase (in saline), while SHAM animals received a striatal injection of 0.9% saline of the same volume. Figure 1 was created with the use of BioRender.com (Toronto, ON)

Experiment 1

The primary endpoint for our first experiment was hematoma volume at 8-hours post-ICH. Rats were randomized into two groups, SAL or HTS, administered once at 2-hours post-ICH, with n = 12 per group before mortality and exclusions (total N = 24). To rule out discomfort due to HTS injections, pain was assessed via the Rat Grimace Scale (RGS), immediately before and after injection.

Experiment 2

The primary endpoint for our second experiment was edema at 24-hours post-ICH, with n = 20 rats per group before mortality and exclusions. Rats were given either SAL or HTS (total N = 40). All animals were assessed with the neurological deficit scale (NDS) at 24-hours post-ICH.

Experiment 3

The primary endpoint for this experiment was cell volume at 24-hours post-ICH, assessed via stereology. Rats received either ICH or SHAM surgeries, with n = 24 ICH rats and n = 12 SHAM rats. These received either SAL or HTS, resulting in 4 groups (n = 12 ICH-SAL rats, n = 12 ICH-HTS rats, n = 6 SHAM-SAL rats, n = 6 SHAM-HTS rats; total N = 36). All animals were assessed with the NDS immediately before euthanasia. Both cell volume and cell density were assessed in ipsilateral CA1 (cornu ammonis 1) and S1 (primary somatosensory cortex). A subset of rats from each group (n = 3 SHAM-SAL rats, n = 3 SHAM-HTS rats, n = 6 ICH-SAL rats and n = 6 ICH-HTS rats) were randomly selected for qualitative analysis of contralateral CA1 subcellular health via TEM, which was assessed in a blinded manner. We limited TEM measurements to a subset of rats for feasibility.

Intracerebral Hemorrhage

ICH surgery was performed as previously described (Rosenberg et al. 1990; MacLellan et al. 2008; Kung et al. 2021). Briefly, rats were anesthetized with isoflurane (4% induction and 2% maintenance, in 70% N₂O and balance O₂) for the duration of the surgery. ICH was induced via bacterial collagenase injection (3µL of Type IV-S, Sigma, 0.2U/µL in saline) into the left striatum (0.5 mm anterior, 3.5 mm lateral, and 6.5 mm depth from bregma (Paxinos and Watson 1986)). Bupivicaine hydrochloride (5 mg/mL, SteriMax, Oakville, ON) was delivered subcutaneously to the site of incision, prior to the cut and prior to closing the wound. A rectal probe was inserted to monitor temperature, which was maintained at ~ 37 °C throughout the surgery using heating mats. Immediately following surgery, all rats were given 5 mL of 0.9% saline subcutaneously, as recommended by our animal care committee for all surgeries causing a large stroke, and allowed to wake up naturally. All rats were given wet Purina rat chow to reduce risk of dehydration and weight loss. SHAM surgeries were induced identically, including all care interventions, but with 0.9% saline injections (rather than collagenase) into the striatum.

Hypertonic Saline Administration

Both SAL and 3% HTS were given at 2 mL/kg and administered via the lateral tail vein under isoflurane anesthesia. Animals were kept under anesthetic for an average of 8 min in Exp. 1 and 6 min in Exps. 2 and 3, in a manner consistent across groups. Tails were gently washed with warm water and placed on a heating mat to facilitate vasodilation. The injection site was cleaned with 70% ethanol prior to injection, and pressure was applied immediately afterwards with gauze to minimize bleeding. Although previous animal work have investigated concentrations as high as 23.4% HTS (Qureshi et al. 1999; Schreibman et al. 2018), clinical studies tend to recommend lower concentrations of HTS, ranging from 3 to 7.5%, partly due to the potential for adverse side effects (e.g., hyperchloremic acidosis) (Fink 2012; Bouchat et al. 2019; Shoamanesh et al. 2021; Holden et al. 2023).

Behavioural Assessments (NDS & RGS)

The Rat Grimace Scale (RGS) was used to assess pain due to injection in Exp. 1 (Sotocina et al. 2011). Rats were filmed immediately before and after HTS or SAL injection to evaluate their facial expressions to quantify pain and discomfort, as previously described (Kung et al. 2021). Orbital tightening, nose and cheek flattening, and whisker and ear changes were assessed on a scale from 0 to 2. Scores were then tallied to produce a composite score ranging from 0 to 10, with 0 indicating no pain and 10 indicating the maximum pain measurable by the scale.

For Exps. 2 and 3, NDS was used to assess functional outcome, as previously described (Kung et al. 2021). Briefly, prior to baseline, rats received one training session to gain familiarity with the specific behavioural tests. Spontaneous circling, beam walking ability, contralateral hindlimb retraction, contralateral forelimb flexion, and bilateral forepaw grasp were assessed for NDS, with the total score ranging from 0 (no deficit) to 14 (severe impairment). A baseline measure was recorded 2 days prior to ICH for comparison with post-ICH scores.

Hemoglobin Assays

To assess the effects of HTS on bleeding, animals were euthanized by decapitation (under isoflurane) 8-hours post-ICH in Exp. 1. Hemoglobin assays were conducted as previously described (Choudhri et al. 1997; Kung et al. 2021). Briefly, extracted brains were separated into ipsilateral hemisphere, contralateral hemisphere, and cerebellum, then homogenized and centrifuged. The supernatant was combined with Drabkin’s reagent and absorbance was recorded at 540 nm. Hematoma volume was calculated by correcting against contralateral blood volume (ipsilateral blood volume – contralateral blood volume = hematoma volume).

Edema

Edema was measured in Exp. 2, at 24-hours post-ICH, as previously described and commonly done (Jiang et al. 2016; Wilkinson et al. 2019; Kung et al. 2021). Briefly, brains were blocked from 2 mm anterior to 4 mm posterior of the collagenase injection site, and then split into ipsilateral striatum and cortex, contralateral striatum and cortex, and cerebellum. Brains were weighed (wet weight), dried at 100 °C for 24 h, and re-weighed (dry weight). Brain water content was determined as .

Tissue Preparation for Histology (Stereology and Transmission Electron Microscopy)

All animals in experiment 3 were euthanized 24 h post-ICH via a ~ 100 mg/kg intraperitoneal injection of sodium pentobarbital (Bimeda MTC, Cambridge ON), as previously described (Kalisvaart et al. 2020). Briefly, rats were perfused with 0.9% saline, followed by Karnovsky’s fixative (Fix and Garman 2000; Kalisvaart et al. 2020). Brains were kept in situ within the skull in fresh fixative for 24 h at 4 °C to prevent histological artefact. After 24 h, brains were split into the ipsilateral hemisphere for stereology, while the contralateral CA1 was dissected out for TEM in 1 × 2 mm blocks.

Stereology

Stereological assessment was conducted as previously described (Wilkinson et al. 2023). Briefly, the ipsilateral hemispheres of brains were sectioned into 80 μm thick coronal sections using a vibratome (Leica VT1200 S), with a random start point to ensure systematic random sampling. Images (three per region) were taken at 40x magnification in ipsilateral medial CA1 (sections ranging from approximately − 3.3 mm to -4.16 mm anterior to bregma) and ipsilateral layer III/IV of S1 (0.7 mm to -0.26 anterior to bregma), using stereotaxic landmarks to remain consistent across animals (Paxinos and Watson 1986). Cell volume and density assessments in ipsilateral CA1 and S1 were made using the nucleator probe and stereological counting rules, respectively, in conjunction with the optical dissector method (Gundersen et al. 1988), as described previously (Wilkinson et al. 2023). An Olympus BX51 microscope outfitted with a XYZ motorized stage controller (Marzhauser Wetzlar, Germany) and calibrated to the Olympus cellSens Dimensions image acquisition and analysis software (Olympus Life Science Solutions, Japan) was used to take these measurements.

Transmission Electron Microscopy

Sample blocks for TEM were first processed (buffer rinsing, secondary fixation, and dehydration), then embedded in resin. Using an ultramicrotome (Reichert-Jung, Ultracut E), semi-thin (1 μm) sections of the contralateral hippocampus were taken and stained with toluidine blue to locate and block off the region of interest (e.g., stratum pyramidale layer of CA1). Ultra-thin sections were taken (90 nm), mounted on grids, and stained with uranyl acetate and lead citrate. Grids were then imaged using a Phillips-FEI Morgagni 268 80 kV (Phillips-FEI, Oregon, USA) transmission electron microscope at several different magnifications by an experimenter blinded to group identity. A minimum of five hundred CA1 neurons were examined per animal, across at least two CA1 tissue sample blocks. Images were then sent to a neuropathologist (FvL) for expert qualitative evaluation of cellular ultrastructure and health across experimental groups, which was conducted in a blinded fashion.

Statistical Analyses

Data were analyzed using GraphPad Prism (v 10.1.0 for Windows, GraphPad Software, Boston, Massachusetts USA). All data are presented as mean ± 95% confidence intervals (CI) for parametric data, and median ± interquartile range (IQR) for non-parametric data. All data were tested for assumptions and analyzed non-parametrically when assumptions were not met. All statistical analyses and procedures were conducted as specified in our a priori planning document (Online Resource 1), except where explicitly noted below. Sample sizes were calculated to provide 80% power for each experiment’s primary endpoint (25% change in hematoma volume for Exp. 1; 1.5% absolute change in edema for Exp. 2; 15% difference in neuronal soma volume for Exp. 3) and increased to account for potential mortality in our large stroke model (~ 10% at 24 h). Detailed justification regarding effect sizes used for all power calculations are provided in our supplemental information (Online Resource 1).

For Exp. 1, hematoma volume was analyzed using a Mann-Whitney test (planned: t-test) due to non-normality. Pain via RGS was analyzed using Mann-Whitney tests. For Exp. 2, edema was compared between groups within brain regions using Mann-Whitney tests and within groups between brain regions using a Friedman test, due to non-normality (planned: two-way ANOVA with Tukey’s HSD). Behaviour in Exps. 1 and 2 were analyzed using Mann-Whitney tests. Additionally, we conducted an unplanned assessment of behaviour across time using a Wilcoxon matched-pairs signed ranks test, and in Exp. 3 analyzed behaviour using a Kruskal-Wallis with Dunn’s multiple comparisons (planned: Mann-Whitney) to allow for comparison across the four experimental groups. We conducted an unplanned compiled behavioural analysis of all animals from Exps. 2 and 3 using a two-way ANOVA with Fisher’s LSD. For Exp. 3, cell density (stereology) was analyzed using two-way ANOVA with Tukey’s HSD. Cell volume (stereology) was analyzed using independent t-tests with Welch’s correction (planned: two-way ANOVA with Tukey’s HSD) due to heteroscedasticity. Subcellular morphology (TEM) were analyzed qualitatively only (planned: two-way ANOVA with Tukey’s HSD). To minimize likelihood of missing a treatment effect, or Type II error, interval and ratio level data were additionally analyzed parametrically, which did not change our findings (data not shown).

Results

Experiment 1

Mortality and Exclusions

One animal from the SAL group was omitted from all analyses due to surgical error. One animal from the HTS group was excluded from RGS analyses due to experimenter error, and one animal from the HTS group was excluded from hematoma volume analyses due to experimenter error. This left final sample sizes of n = 11 SAL and n = 12 HTS for RGS analyses, and n = 11 per group for hematoma volume analyses. All relevant raw data are provided in our supplemental information (Online Resource 2).

Pain

Although no pre-ICH baseline was taken, pre-dose RGS scores (median 3.86, [3.74, 3.97] IQR) were well above normal levels (e.g., 0–1), as expected following an ICH. No differences between HTS and SAL groups were observed, either pre- (Fig. 2a; p = 0.0966) or post-dose (p = 0.7597). Thus, injection of HTS did not appear to cause any additional pain in these animals.

Fig. 2.

Experimental Results In Exp. 1, pain (a) was not significantly different between 3% hypertonic saline (HTS) and 0.9% saline (SAL) groups immediately pre-dose (i.e., 1.5 h post-stroke) (p = 0.0966) or immediately post-dose (i.e., 2.5 h post-stroke) (p = 0.7597). Hematoma volume (b) also did not significantly differ between groups (p = 0.6522). In Exp. 2, no differences were seen in either NDS (c; p = 0.1320) or edema (d; p > 0.4797) between groups. In Exp. 3, NDS (e) was significantly different between SHAM and ICH groups (p < 0.0266) but no differences were observed between SAL and HTS groups (p > 0.9999). NDS scores from ICH animals were compiled across Exps. 2 & 3 (f), and there were no significant differences between HTS and SAL groups (p = 0.6479). IPSI HEM = Ipsilateral hemisphere, CONTRA HEM = Contralateral hemisphere, IC = Ipsilateral Cortex, CC = Contralateral Cortex, IS = Ipsilateral Striatum, CS = Contralateral Striatum, Cb = Cerebellum

Hematoma Volume

No differences in hematoma volume were found between HTS or SAL groups 8 h post-ICH (Fig. 2b; p = 0.6522). Generally, the bleed volumes were substantial (108.2 µL median, [92.5, 126.6] IQR).

Experiment 2

Mortality and Exclusions

One animal from the SAL group was excluded from the entire experiment due to surgical error. Two additional animals were excluded from all analyses, one from the HTS group due to a spontaneous death, and one from the SAL group was euthanized early due to high morbidity. This left final group sizes of n = 19 HTS rats and n = 18 SAL rats.

Behaviour

ICH significantly worsened neurological deficit scores (p < 0.0001). No differences in NDS scores were found between 3% HTS and 0.9% SAL treated groups 24 h post-ICH (Fig. 2c; p = 0.1320). Thus, HTS did not influence neurological deficit scores.

Edema

Brain water content was significantly elevated in ipsilateral striatum and cortex (versus contralateral side) for both groups (p ≤ 0.0121) at 24 h post-stroke. There was no significant effect of HTS in any region (Fig. 2d), including ipsilateral cortex (p = 0.4797), contralateral cortex (p = 0.6415), ipsilateral striatum (p = 0.8453), or contralateral striatum (p = 0.9880). As expected, brain water content in the cerebellum was not elevated (median 77.9%, [77.7, 78.1] IQR) and did not differ between groups (p = 0.5378). Thus, ICH caused significant edema that was not mitigated by HTS treatment.

Experiment 3

Mortality and Exclusions

Three animals were excluded from all analyses: two ICH-HTS animals were euthanized early over humane concerns, and one ICH-SAL animal spontaneously died. This left group sizes of n = 11 ICH-SAL rats, n = 10 ICH-HTS rats, n = 6 SHAM-SAL rats, n = 6 SHAM-HTS rats for analyses.

Some animals were further excluded from stereology due to histological artefact, determined by a blinded researcher, via standards defined previously (Kalisvaart et al. 2020). Specifically, two ICH-HTS animals were excluded from S1 analyses. One ICH-HTS, 1 SHAM-SAL, and 1 SHAM-HTS animal were excluded from CA1 analyses. There were no exclusions in TEM groups.

Behaviour

Behaviour was significantly different among our four groups (Fig. 2e; p < 0.0001), which was driven primarily by differences between SHAM-SAL and ICH-SAL (p = 0.0004), and SHAM-HTS and ICH-HTS (p = 0.0266) groups. No significant differences between SAL and HTS groups were found (p > 0.9999).

When behaviour was compiled across Exps. 2 and 3, ICH-HTS and ICH-SAL groups also did not significantly differ (Fig. 2f; p = 0.6479). Thus, there was no evidence for HTS reducing the neurological deficit caused by ICH.

Ipsilateral Cell Volume and Density

Cell soma volume did not differ between any groups for CA1 (Fig. 3a) or S1 (Fig. 3b). For ICH groups, HTS did not affect cell volume in CA1 (p = 0.1461) or S1 (p = 0.3519). For HTS groups, ICH did not affect cell volume in CA1 (p = 0.3657) or S1 (p = 0.4028). For SAL, ICH did not affect cell volume in CA1 (p = 0.3973) or S1 (p = 0.3656). ICH had no effect on cell density counts, in either CA1 (Fig. 3c; p = 0.7923) or S1 (Fig. 3d; p = 0.4219); likewise, HTS had no effect on CA1 cell counts (p = 0.0983) or S1 cell counts (p = 0.5741).

Fig. 3.

Cell Volume and Density Results Neither ICH (p = 0.3657) nor HTS (p = 0.1461) affected CA1 cell volume (a). Neither ICH (p = 0.4028) nor HTS (p = 0.3519) affected S1 cell volume (b). Similarly, ICH (p = 0.7923) and HTS (p = 0.0983) did not affect CA1 cell counts (c); ICH (p = 0.4219) and HTS (p = 0.5741) did not affect S1 cell counts (d) either

Contralateral Cellular Health

Following administration of either HTS or SAL, all SHAM animals displayed ultrastructural signs of osmotic stress within contralateral CA1 (Fig. 4a-h). In SHAM-SAL animals (Fig. 4a-d), cellular morphology predominantly remained intact, though the soma of some pyramidal neurons contained edematous organelles (e.g., Golgi apparatus, endoplasmic reticulum) and evidence of mitochondrial injury/stress (e.g., disorganized cristae, signs of lysosomal transition). Axons also displayed signs of mild osmotic stress, such as edema, decompaction or thinning of myelin, and peri-axonal dilation. Ultrastructural signs of osmotic stress were pronounced in SHAM-HTS animals (Fig. 4e-h), with disrupted axonal integrity and myelination (e.g., membranous whorls, electron dense “smudging”), and high frequency of lysosomal bodies as well as edematous organelles. In both SHAM-SAL and SHAM-HTS groups (especially so in SHAM-HTS animals), plasma and nuclear membranes of CA1 pyramidal neurons appeared crenated, with a high incidence of intranuclear vacuoles and invaginations (Fig. 4b, e, f); interestingly, this is morphology that we have previously associated with reduced cell volume in area CA1 (Kalisvaart et al. 2020). Taken together, the morphological changes we observed in SHAM groups were especially pronounced in proximity to blood vessels (Fig. 4e), and were reminiscent of ODS, in which a rapid overcorrection of systemic hyponatremia via administration of HTS results in cellular injury (Bouchat et al. 2019). The pathology observed in SHAM-SAL animals is distinct from that of animals who display ultrastructural evidence of perfusion and fixation artefact, and is not observed in historical shams who only received a standard subcutaneous injection of 0.9% saline for hydration at the time of their procedure (Fig. 5a-i) (Kalisvaart et al. 2020). Here, in addition to subcutaneous hydration, SHAM-SAL rats received 0.9% saline infusions into striatum (SHAM stroke – no ICH), as well as the tail vein (control treatment to compare to the 3% HTS group, administered twice, Fig. 1). In comparison, our historical shams shown in Fig. 5d-f (Kalisvaart et al. 2020) did not receive repeated venous injections of 0.9% saline, nor did they receive striatal infusions of 0.9% saline. It is very unlikely that the striatal injection of 0.9% saline (i.e., differences in sham surgery methodology) account for our ultrastructural observations in SHAM-SAL animals, given that these effects are observed within distal contralateral regions far removed from the site of striatal injection. Additionally, striatal saline injections were minute (3 µL) and given a full 24 h prior to euthanasia. Instead, the more parsimonious explanation is that repeated bolus administration of 0.9% saline by venous injection explain the ultrastructural effects observed in SHAM-SAL animals Fig. 5g-i.

Fig. 4.

Comparison of Hippocampal Ultrastructural Morphology in Sham and ICH Groups Treated with Either 0.9% or 3% Saline in Experiment 3. Representative TEM images depict layer CA1 of the hippocampus in the contralateral (right) hemisphere 24 h post-procedure in SHAM groups treated with SAL (a, b, c, d) or HTS (e, f, g, h), as well as ICH groups treated with SAL (i, j, k, l) or HTS (m, n, o, p). Both SHAM groups display evidence of osmotic stress, evidenced by edematous changes to organelles, formation of intranuclear vacuoles and crenated plasma membranes (some examples denoted by #), and loss of myelin integrity (denoted by solid arrows). Mitochondria either became hypertrophic and elongated (denoted by stars; a, b, e, f, g, h), or became electron dense and underwent lysosomal transition (denoted by *; a, b, d, e, f, h), while in some areas, rough endoplasmic reticulum became dilated with a high ribosomal density (denoted by arrow heads; d, e, f, h), directly indicative of osmotic stress. Myelin sheaths surrounding axons appeared electron dense, with a loss of lamellar organization (denoted by solid arrows; a, c, d, e, f, g). These changes were pronounced in SHAM-HTS animals, especially in proximity to blood vessels (denoted “BV”, e). In ICH groups, signs of ICH related injury were prevalent, such as cells undergoing degenerative changes, evidenced by either edematous mitochondria (denoted by diamonds; j, l, m, n, o, p) or electron dense mitochondria (denoted by *; i, j, l, m, n, p), edematous endoplasmic reticulum (denoted by arrow heads in j, p), and loss of axon and myelin integrity (denoted by solid arrows; j, k, l, m, n, o) accompanied by membranous whorls (denoted by open arrows; k). In some cases, formation of phagocytic vacuoles for microglial digestion of dysfunctional organelles was observed (i). In ICH-SAL animals, some pyramidal neurons appeared shrunken, dense, and crenated (top neuron; j), while others had an edematous cytoplasm (bottom neuron; j). In ICH-HTS animals, pyramidal neurons were extremely vacuolated and edematous in appearance (m, p), with pronounced interlamellar distension and splitting of myelinated axons (m, o). This pathology was especially pronounced in proximity to blood vessels, which displayed degenerative changes to pericytes and endothelial cells, surrounded by edematous astrocytic processes and neuropil (n). Scale bars are shown in the lower lefthand corner of each image

Fig. 5.

Comparison of hippocampal ultrastructural morphology in 0.9% Saline Treated Shams versus Artefactual and Historical Data. Representative TEM images depict layer CA1 of the hippocampus in sham animals inadequately perfused and fixed (a-c), in historical sham tissue with appropriate fixation (d-f) (unpublished images from Kalisvaart et al. 2020), and SHAM-SAL animals in the present study (g-i), all at 24 h post-sham procedure. Historical shams did not receive repeated venous administration of 0.9% saline, while SHAM-SAL animals did. Characteristic morphology of perfusion and fixation artefact includes presence of “dark neurons” with an electron dense cytoplasm and nucleoplasm (a) along with widespread edematous changes to organelles regardless of cell type (a, b, c); mitochondria (denoted by *) become bloated and lose organization of cristae, or begin to undergo lysosomal transition, while the tubular and lamellar organization of endoplasmic reticulum (denoted by arrow heads) is lost (a, b). The myelin sheath (denoted by solid arrows) itself remains predominantly intact, however, despite edematous changes to the axolemma and the organelles it contains (a, c). Comparatively, neurons in historical shams are healthy in appearance, with distinct cytoplasmic and nucleoplasmic compartments, and structurally healthy organelles (d, e, f). Axons appear normal, with predominantly compact lamellar myelin organization (d, e, f). In contrast, tissue from SHAM-SAL animals collected for the present study depicts unique signs of osmotic stress not typical of either historical shams or those with perfusion/fixation artefact (g, h, i). A greater proportion of mitochondria appear in lysosomal transition, and remaining mitochondria are hypertrophic and elongated, with some cristae disorganization (g, h, i). Additionally, ribosomal density surrounding dilated rough endoplasmic reticulum is increased, a characteristic sign of osmotic stress (g, h). Myelinated axons lost myelin sheath integrity, with lamellar splitting, formation of membranous whorls, and electron dense changes (g, i). Scale bars are shown in the lower lefthand corner of each image

Signs of additional ICH-related injury were present in CA1 of the contralateral hemisphere, such as scattered degenerating cells (e.g., neurons and oligodendrocytes), degenerating and edematous organelles, elongated mitochondria, and loss of axonal/myelin integrity in both ICH-HTS and ICH-SAL animals (Fig. 4i-p). The severity of pathology in ICH-SAL animals was more pronounced compared to SHAM-SAL animals, as expected. This contralateral hippocampal cellular injury was not simply due to extension of the ipsilateral striatal bleed, and therefore more likely reflects compromised microvascular perfusion due to the compressive effects of elevated ICP. We observed that ICH-SAL animals (Fig. 4i-l) featured some pyramidal neurons that appeared shrunken, dense, and crenated, while other neurons featured a swollen and edematous cytoplasm. Conversely, in ICH-HTS animals (Fig. 4m-p), the majority of CA1 pyramidal neurons observed had an edematous and vacuolated cytoplasm, often bloated in appearance, with pronounced interlamellar distension and splitting in myelinated axons; yet the nucleolus and cytoplasmic membrane of these cells remained intact. Endothelial cells and pericytes surrounding blood vessels in ICH-HTS animals displayed signs of degeneration and early necrosis, and associated astrocytic endfeet and processes were similarly edematous as compared to surrounding neuronal structures within the neuropil (Fig. 4n).

Discussion

Overall, our results suggest a lack of benefit of 3% HTS administration following a large striatal bleed. As expected, HTS did not impact bleeding, with no differences in hematoma volume observed between groups at 8-hours post-ICH. However, HTS administration did not ameliorate edema or behavioural deficits at 24-hours post-ICH, a time at which there is extensive cytotoxic and ionic edema (Simard et al. 2007; Ironside et al. 2019). On the ultrastructural level, we did find qualitative evidence that systemic HTS administration exacerbates cellular stress in regions distal to the bleed, such as contralateral hippocampus. Despite these ultrastructural perturbations, we did not find any evidence of tissue compliance after ICH, an effect we have observed previously following large stroke (Kalisvaart et al. 2020; Wilkinson et al. 2023). Taken together, our data suggest that HTS does not improve outcome after ICH in rats, and may even detrimentally impact subcellular health, at least under this dosing regimen. Therefore, our findings indicate the urgent need for further research on how HTS may affect cellular health.

We found that HTS did not reduce edema or improve functional outcome after ICH. While this conflicts with previous preclinical studies (Schreibman et al. 2018), our data are in line with clinical findings and conclusions (Shah et al. 2018; Shoamanesh et al. 2021; Chugh et al. 2021; Barik et al. 2023). Owing to numerous differences among studies (e.g., bleed size, animal species and strain, stroke model, post-operative animal care, etc.), it is difficult to identify a definitive factor to explain these discrepant findings. However, our bleeds were notably large (median 108.2 µL bleed). Scaling up based on brain weights, assuming an average rat brain weight of 2 g and an average human brain weight of 1.4 kg (Piao et al. 2013), our bleeds are roughly comparable to a ~ 75 mL bleeds in humans. Large bleeds result in abundant serum extrusion as clot retraction occurs, coinciding with the development of extensive vasogenic edema due to elevated thrombin levels, neuroinflammation, and BBB injury in perihematomal regions (Ironside et al. 2019; Chen et al. 2021). In addition to creating a large bleed, the collagenase model is also known to model extensive BBB injury (MacLellan et al. 2008; Nadeau et al. 2019; Jia et al. 2021). Therefore, HTS alone, which only reduces cytotoxic and ionic edema, may be insufficient to reduce brain water content in this setting. While we could have used milder bleeds, our large bleeds presumably have better translational relevance, as decompensated ICP tends to occur in the setting of large strokes (Wilkinson et al. 2020). Another possibility is that HTS may contribute to faster edema resolution, which would require longer survival times to test; for humane reasons, our latest endpoint was set at 24 h post-ICH. This may have influenced our results, as edema peaks on day 3 post-stroke in the collagenase model (Ironside et al. 2019; Jiang et al. 2022). However, given that cytotoxic and ionic edema peak early (Ironside et al. 2019; Chen et al. 2021), the survival times used in our experiment should have permitted us to observe any potential benefit that could be attained from HTS administration. It remains possible that 3% HTS could have transient benefits on edema or other outcomes, an effect that may have dissipated between the last dose of 3% HTS (14 h post-stroke) and outcome assessment (24 h post-stroke). Thus, this intervention may simply fail to afford lasting benefit, at least with the doses and concentrations used in this study.

Following ICH, 3% HTS administration paradoxically resulted in dramatic neuronal and astrocytic swelling in distal uninjured regions (e.g., contralateral hippocampus). In ICH-SAL animals, this effect was less pronounced, with some pyramidal neurons displaying edematous changes while others appeared crenated and shrunken. The latter is more typical of the “tissue compliance” morphology we have previously observed in ICH animals, which occurs in the absence of contralateral hippocampal edema (Kalisvaart et al. 2020; Wilkinson et al. 2023). Comparatively, in our SHAM (no-stroke) animals, HTS administration had adverse effects on cellular health in a manner characteristic of osmotic injury, with morphology reminiscent of rodent ODS models (Bouchat et al. 2019; Gilloteaux et al. 2020). A similar but milder effect was observed in SHAM-SAL animals. As ODS occurs due to an rapid correction or overcorrection of hyponatremia, our dosing regimen or administration route could potentially account for our findings (Martin 2004). In both SHAM groups, CA1 pyramidal neurons displayed intranuclear vacuoles, invaginations, and crenated membranes (morphology typically associated with cell volume reduction (Kalisvaart et al. 2020). Overall, the differential effects of saline on cellular morphology in our groups may have ultimately masked the tissue compliance effect (e.g., cellular shrinkage in SHAM-HTS, cellular swelling in ICH-HTS). To indirectly investigate this, we compared our SHAM stereology data (e.g., cell volume and density) to that of historical SHAMs (Kalisvaart et al. 2020), and observed trends that may support this hypothesis (Online Resource 3). However, comparisons to historical controls are not ideal, and further research is needed to investigate this directly.

Despite subcellular perturbations across groups, we did not observe any corresponding behavioural differences. Perhaps more sensitive tests of functional outcome (MacLellan et al. 2006) may have been able to identify some subtle effects of HTS administration. Alternatively, HTS administration may reduce edema in perihematomal regions and spare proximal neurons, but this functional benefit could be inadvertently masked by possible adverse effects of HTS on more distal regions. Future studies should implement a battery of behavioural tests or alternative methods to elucidate regional functional impairments, such as electrophysiology, to more sensitively assess such nuances over the acute post-stroke period. If administration of 3% HTS does acutely exacerbate cellular stress in brain regions distal to the bleed, assessments of cellular health and corresponding functional and behavioural outcomes should also be conducted over the long term to determine whether there are any lasting consequences of this effect, although this may pose some challenges when using a model of large ICH. Lastly, future studies should consider employing quantitative TEM methods to better identify the extent of subcellular perturbation observed with HTS administration.

As with any negative finding of treatment efficacy, one must consider dosage, timing, and choice of endpoints. Specific to HTS, the lack of consensus throughout osmotic therapy literature regarding dosage, regimen, and timing may explain why some studies have found benefit and we have not. For example, HTS concentrations as low as 3% and as high as 23.4% are administered both continuously and intermittently in clinic (Wagner et al. 2011; Fink 2012; Barik et al. 2023; Donepudi et al. 2024; Khasiyev et al. 2024). With such heterogeneity in the literature, treatment decisions are largely left to physician preference (Schrot and Muizelaar 2005; Tyagi et al. 2007; Fink 2012), making it difficult to compare across studies. Previous preclinical ICH studies have found benefit with administration of 23.4% HTS (Qureshi et al. 1999; Schreibman et al. 2018), while our study opted for a regimen we deemed to be safer and more clinically relevant: 3% HTS, administered intermittently and intravenously. Perhaps higher concentration of HTS, more frequent dosing, or an alternative administration route may have improved outcomes after ICH. However, 3% HTS is commonly used and/or advised in clinic (Wagner et al. 2011; Shoamanesh et al. 2021; Jiang et al. 2022), and higher doses of HTS (such as 23.4%) may lead to unacceptable consequences to subcellular health. Dose-response work on HTS in the setting of both preclinical and clinical ICH is urgently needed, especially given the potential impact on cellular health. Eventual synthesis of these studies through systematic and meta-analytic means will provide a clearer perspective on the true efficacy (or lack thereof) of HTS in pre-clinical ICH settings. This would better inform future clinical investigations, but as it currently stands, there is not sufficient literature in this area to conduct such an analysis.

Our study has several limitations. Due to the desire to avoid confounds, we did not measure ICP in our animals, which prevented us from assessing the effects of HTS on global ICP levels. Future studies should directly assess ICP and subcellular health to identify any potential associations between ICP, tissue compliance, and subcellular health. We did not record water intake or measure plasma osmolarity in our study to avoid the stress of repeated blood sampling, which precluded us from relating changes in serum osmolarity to our cellular and subcellular findings. Lastly, we acknowledge that our ultrastructural findings were qualitative in nature but emphasize that this evidence fits with well-known ODS findings and thus warrants further translational study.

In conclusion, we did not observe obvious benefit of HTS on behavioural outcomes or edema in the setting of large experimental striatal ICH, and rather noted preliminary evidence of treatment-related cellular stress in regions distal to the bleed. Future studies evaluating the utility of HTS as an ICH treatment should evaluate the impact on cellular health across both edematous and non-edematous regions using a variety of dosing regimens and additional ICH models. Overall, further research is urgently needed to more fully characterize the effect of HTS administration.

Electronic Supplementary Material

Below is the link to the electronic supplementary material.

Author Contributions

TK, AK, and FC conceptualized the study with input from GJ and FvL. TK, AK and AS collected data, which was reviewed by FvL (TEM images) and FC (all data). TK and AK drafted the manuscript with oversight from FC and input from others. All authors approved the manuscript. FC was project and student supervisor.

Funding

Research supported by the Canadian Institutes of Health Research (CIHR, project grant #174992) to FC (PI), AK, FvL, GJ and others. TK and AK were supported by CIHR Canada Graduate Scholarship Doctoral awards. AK was also supported by an Izaak Walton Killam Memorial Scholarship.

Data Availability

Numerical data (used for statistical analyses) are provided in a supplementary file.

Declarations

Competing Interests

The authors declare no competing interests.