All experimental procedures were performed after approval from the Animal Welfare Body of the Charles University in Prague, Faculty of Medicine in Hradec Kralove, Czech Republic (approval no. 17-16/2014-6848) in accordance with Czech legislation on the protection of animals, which complies with Directive 2010/63/EU of the European Parliament and Council. Ten male and female rabbits (New Zealand white rabbit; weight, 2.0–3.0 kg; VELAZ 34081/2008-10001, CZ 21906828, Únětice, Czech Republic) were included in the study. The animals were housed in a standard cage at 21 °C under a 12-h dark/12-h light cycle with unrestricted access to laboratory chow and tap water. After a 1-week acclimatization period, the rabbits were used for the study.
Anesthesia and surgical preparation
After an overnight fast with unrestricted access to tap water, the rabbits were anesthetized using an intramuscular induction dose of ketamine (40 mg/kg) and xylazine (4 mg/kg). The animals were placed in the supine position on an operating table. The body locations used for cannulation, electrocardiogram electrodes, and tracheostomy were shaved. Intravascular cannulas (Vasofix® Safety, B.Braun, Melsungen, Germany) were inserted in both marginal ear veins (G24) and the right central ear artery (G22) for continuous blood pressure monitoring, arterial blood gas analysis, and continuous infusion of a balanced crystaloid solution (Ringerfundin, B. Braun, Melsungen, Germany, 10 ml/kg/h), anesthesia, and a muscle relaxant. Mean arterial blood pressure was maintained above 50 mmHg with norepinephrine infusion as necessary.
The animals were tracheotomized after they were stabilized hemodynamically. A cuffless tracheal tube with an outer diameter of 2.5 mm was inserted between the third and fourth tracheal rings. After verifying correct placement by auscultation, mechanical ventilation was initiated using an anesthesia machine (Cirrus Trans2/Vent 2, Datex, Helsinki, Finland) with initial settings of pressure-controlled ventilation, respiratory rate of 40 breaths/min, inspiratory pressure of 14–16 cm H2O according to the weight of the rabbit, and a positive end-expiratory pressure of 3 cm H2O (lowest value on the ventilator), which was adjusted according to the first blood gas analysis results. The ventilator setting was not changed after osmotherapy. Mean arterial blood pressure (MAP), heart rate, and rectal temperature were recorded throughout the study. Rectal temperature was maintained at 38.5–39.5 °C using a heating plate and a thermoisolation blanket. Balanced anesthesia was maintained using isoflurane (0.6–1 vol%, Forane, AbbVie Inc., Chicago, IL, USA) in a mixture of 1 l/min oxygen and 1.2 l air with an inspiratory oxygen fraction (FiO2) of 50–55 %, continuous intravenous infusion of fentanyl (0.4 μg/kg/min, Fentanyl Torrex, Chiesi Pharmaceuticals GmbH, Vienna, Austria), and the muscle relaxant pipecuronium bromide (0.6 mg/kg/h, Arduan, Gedeon Richter Plc., Budapest, Hungary).
Each animal was subsequently rotated into the prone position, and the right temporo-parieto-occipital area of the head was shaved. The skin and periosteum of the skull were incised and reflected, and bleeding was stopped by bipolar electrocoagulation. The margins of the exposed area were determined by the midline, the base of the right ear, the external occipital protuberance, and the right caudal supraorbital process. A 3-mm hole was drilled through the exposed skull and was increased in size using a mosquito pean. The final size of the cranial window was obtained using a Kerrisson rongeur. Bleeding from the diploe was stopped using bone wax. The dura mater was cut carefully around the edges of the cranial window using microscissors to minimize brain surface injury. The dimensions of the cranial window were approximately 12 × 8 mm, with intact arachnoid mater at the base of the window. A 15-min stabilization period was maintained after controlling bleeding. During this period, the wound was flushed frequently with sterile 37 °C normal saline, hemodynamic data were recorded, a sample of arterial blood was sent for laboratory examination (levels of blood gases, sodium, potassium, chlorides, and hemoglobin), a SDF probe was attached to the brain surface, and initial SDF imaging was performed.
SDF imaging procedure
The SDF imaging probe was covered with a sterile plastic sheath and placed above the target tissue; a conventional hand-held technique was used. The sites of interest on the brain surface were selected randomly. Exposed tissues, other than those covered by the SDF imaging probe, were moisturized intermittently using 37 °C sterile normal saline. SDF imaging data were recorded digitally from three different areas (fields) within the site of interest for each animal at each measurement, and video clips lasting at least 20 s were recorded from each area (total of three video clips). Analysis of flow in larger vessels was used as a quality control measure to ensure that excessive pressure was not applied to the tissue .
Two animals were used to test the feasibility of this model, and 18 animals were included in the study. Two animals died during the instrumentation phase. Sixteen animals were randomized (a computer-generated random list of animals was used) to receive 3.75 ml/kg body weight of either 3.2 % HTS (HTS group) or 20 % mannitol (MTL group) solution administered intravenously over 15 min using an infusion pump after the initial SDF measurement. Both solutions had the same osmolarity (1099 mOsm/l) and were infused via a peripheral venous catheter. The 3.2 % HTS solution was prepared by the hospital pharmacy. The volume infused was equivalent to a dose of 0.75 g/kg body weight MTL. A second set of SDF imaging data were obtained, hemodynamic data were recorded, and a sample of arterial blood was sent for laboratory examination (including blood lactate levels) 20 min after infusing MTL or HTS. The animals were sacrificed at the end of the experiment using an overdose of thiopentone (30 mg/kg body weight).
The team members were blinded to the assigned groups while preparing the animals, acquiring data, analyzing the video clips, and conducting the statistical analysis. The administered solution was drawn up off-site and administered by an unblinded co-worker (a laboratory staff member).
Video clips were randomly coded and analyzed offline by a single observer blinded to file order. Two clearest and most stable parts of each video clip (sequences) that met the software’s stability criteria were selected for the analysis. Flow in larger vessels was checked to ensure that excessive pressure was not applied during recording. A total of six sequences were analyzed per animal per measurement, and the average was used for subsequent calculations. The final on-screen magnification of the images obtained using the SDF imaging device was 325-fold the original, and the actual size of the field evaluated was 1280 × 960 μm.
Microcirculatory parameters were measured using AVA V3.0 software (AMC, University of Amsterdam, Netherlands). To decrease possible inter-observer variability, all analyses were performed by a single, blinded researcher (VDjr).
The following parameters were analyzed offline:
Total small-vessel density (SVD) and all-vessel density (TVD) were defined as the total length of the respective vessels inside the image divided by the total area of the image. Small vessels were defined as those with diameters ≤ 25 μm .
The DeBacker score, given in mm−1, was defined as the number of vessels crossing three arbitrary horizontal and three vertical equidistant lines (drawn on the screen) divided by the total length of the lines .
Microvascular flow index was calculated as an average value of the semiquantitative score (0 = absent flow, 1 = intermittent flow, 2 = continuous sluggish [slow] flow, 3 = continuous [normal] flow, 4 = hyperdynamic [fast] flow) of the microvascular flow in the four image quadrants, as assessed subjectively by an observer . Absent flow was defined as no flow throughout the recorded image, intermittent flow as at least 50 % of the recorded time with no flow, sluggish flow as continuous but slow flow, and continuous flow as fast flow lasting throughout the recording.
The proportion of perfused vessels (PPV) was defined as the percentage of all visible vessels with at least sluggish flow. Perfused small vessel density (PSVD) and perfused vessel density (PVD) were obtained as SVD and TVD multiplied by the respective proportion of perfused vessels.
A power analysis using an α error of 0.05 and β error of 0.2 was performed based on previously published brain microcirculatory data for rabbits  using MedCalc 7.6.0. (MedCalc Software, Ostend, Belgium). The sample size needed for the t-test (independent groups) to detect a 15 % difference in SVD or TVD and a 20 % difference in the DeBacker score was calculated. This calculation yielded a sample size of 16 subjects (eight subjects per group). The sample size was increased to 18 animals to compensate for potential dropouts and inaccurate predictions used for the power analysis.
The continuous variable results are presented as means ± standard deviations or as medians with interquartile ranges based on the results of a test of the normality of the distribution using a one-sample Kolmogorov-Smirnov test. Differences in sex were analyzed using Fisher’s exact test. The Mann–Whitney U-test was used to compare results between groups when the sample distribution was not normal (pH, PaCO2, PPV, MFI, dose of catecholamines, and lactate levels). An unpaired t-test was used to compare all other results between groups. A P value < 0.05 was considered to indicate significance. The statistical analysis was performed using MedCalc 7.6.0.