Evaluation of Cerebral Blood Flow Autoregulation in the Rat Using Laser Doppler Flowmetry

Summary

This article demonstrates the use of laser Doppler flowmetry to evaluate the ability of the cerebral circulation to autoregulate its blood flow during reductions in arterial blood pressure.

Abstract

When investigating the body's mechanisms for regulating cerebral blood flow, a relative measurement of microcirculatory blood flow can be obtained using laser Doppler flowmetry (LDF). This paper demonstrates a closed skull preparation that allows cerebral blood flow to be assessed without penetrating the skull or installing a chamber or cerebral window. To evaluate autoregulatory mechanisms, a model of controlled blood pressure reduction via graded hemorrhage can be utilized while simultaneously employing LDF. This enables the real time tracking of the relative changes in the blood flow in response to reductions in arterial blood pressure produced by the withdrawal of circulating blood volume. This paradigm is a valuable approach to study cerebral blood flow autoregulation during reductions in arterial blood pressure and, with minor modifications in the protocol, is also valuable as an experimental model of hemorrhagic shock. In addition to evaluating autoregulatory responses, LDF can be used to monitor the cortical blood flow when investigating metabolic, myogenic, endothelial, humoral, or neural mechanisms that regulate cerebral blood flow and the impact of various experimental interventions and pathological conditions on cerebral blood flow.

Introduction

Autoregulatory mechanisms in the cerebral circulation play a crucial role in maintaining homeostasis and normal function in the brain. Autoregulation of the cerebral blood flow is affected by multiple factors including heart rate, blood velocity, perfusion pressure, the diameter of the cerebral resistance arteries, and the microcirculatory resistance, all of which play a role in maintaining the total cerebral blood flow constant in the brain over the physiological range of systemic blood pressures. When arterial pressure increases, these mechanisms constrict arterioles and resistance arteries to prevent dangerous increases in intracranial pressure. When arterial blood pressure decreases, local control mechanisms dilate the arterioles to maintain tissue perfusion and O₂ delivery. Various pathological conditions such as hypercapnia, traumatic or global hypoxic brain injury, and diabetic microangiopathy^1,2,3,4,5,6 may disrupt the brain's ability to autoregulate its blood flow. For example, chronic hypertension shifts the effective autoregulatory range toward higher pressures^7,8,9, and a high salt (HS) diet not only interferes with normal endothelium-dependent dilation in the cerebral microcirculation¹⁰, but also impairs the ability of autoregulatory mechanisms in the cerebral circulation to dilate and maintain tissue perfusion when arterial pressure is reduced¹¹. Cerebral autoregulation is also impaired in Dahl salt-sensitive rats when they are fed a HS diet¹².

During reductions in arterial pressure, dilation of the cerebral resistance arteries and arterioles initially returns cerebral blood flow to control values despite the reduced perfusion pressure. As arterial pressure is reduced further, cerebral blood flow remains constant at the lower pressure (plateau phase of the autoregulatory response) until the vasculature can no longer dilate to maintain blood flow at the lower pressure. The lowest pressure at which an organ can maintain normal blood flow is termed the lower limit of autoregulation (LLA). At pressures below the LLA, cerebral blood flow decreases significantly from resting values and decreases in a linear fashion with each reduction in arterial perfusion pressure^13,14. An upward shift in the LLA, as observed in hypertension^7,8,9, may increase the risk and severity of ischemic injury during conditions where the arterial perfusion pressure is reduced (e.g., myocardial infarction, ischemic stroke, or circulatory shock).

LDF has proven to be an extremely valuable approach to evaluate the blood flow in the microcirculation under a variety of circumstances, including autoregulation of the blood flow in the cerebral circulation^11,14,15. In addition to evaluating autoregulatory responses, LDF can be used to monitor the cortical blood flow when investigating metabolic, myogenic, endothelial, humoral, or neural mechanisms that regulate the cerebral blood flow and the impact of various experimental interventions and pathological conditions on cerebral blood flow^{10,16,17,18,19,20,21}.

LDF measures the shift in reflected laser light in response to the number and velocity of moving particles--in this case, red blood cells (RBC). For studies of cerebral vascular autoregulation, arterial blood pressure is changed either by the infusion of an alpha-adrenergic agonist to increase arterial pressure (because the cerebral circulation itself is insensitive to alpha-adrenergic vasoconstrictor agonists)^12,15 or via controlled blood volume withdrawal to reduce arterial pressure^11,14. In the present study, LDF is utilized to demonstrate the effects of graded reductions in blood pressure on cerebral autoregulation in a healthy rat. Although open and closed skull methods have been described in the literature^22,23,24,25, the present paper demonstrates a closed skull preparation, allowing cerebral blood flow to be assessed without penetrating the skull or installing a chamber or cerebral window.

Protocol

The Medical College of Wisconsin Institutional Animal Care and Use Committee (IACUC) approved all protocols described in this paper and all procedures are in compliance with the National Institutes of Health (NIH) Office of Laboratory Animal Welfare (OLAW) regulations.

1. Experimental animals and preparation for recording

1. Use 8–12-week-old male Sprague-Dawley rats weighing 250–300 g. For these experiments, feed rats a standard diet consisting of 0.4% NaCl, 200 g/kg casein, 3 g/kg DL-methionine, 497.77 g/kg sucrose, 150 g/kg cornstarch, 50 g/kg corn oil, 50 g/kg cellulose, 2 g/kg choline bitartrate, 35 g/kg mineral mix, and 10 g/kg vitamin mix.
2. Record arterial blood pressure and LDF readings using data acquisition software or any comparable recording method.
3. Attach the arterial pressure transducer to one channel of the recording system and the LDF probe to the other channel on the recording system.
4. Prior to the measurement, calibrate the laser Doppler probe to set a motility standard and ensure that the laser Doppler flowmeter is providing a steady output.
5. Prepare additional equipment needed for the preparatory surgery and for the experiment: a dissecting microscope, a rodent ventilator, an end tidal CO₂ monitor, a stereotaxic instrument to fix the rat's head in position, and a micromanipulator to locate the LDF probe over the pial microcirculation and maintain it in a steady position.

2. Surgical preparation

1. Weigh the rat and anesthetize the animal in an induction chamber with 3.5% isoflurane and 30% O₂ supplement.
2. Remove the animal from the induction chamber and substitute an anesthetic mask delivering 1.5–3% isoflurane with a 30% O₂ supplement.
3. Place the rat on a circulating water blanket maintained at 37 °C and check reflexes with a toe pinch to ensure that there is a withdrawal reflex. Apply sterile ophthalmic ointment to both eyes to prevent corneal desiccation.
4. Shave the top of the cranium, ventral neck area, and femoral triangles. Remove any loose hair from those areas and clean with rubbing alcohol.
5. Place the rat in a supine position on a heating pad with a circulating warm water pump to maintain the animal's body temperature at 37 °C and temporarily secure it to the pad using medical tape.
6. Install a tracheal cannula (PE240 polyethylene tubing) through a ventral incision in the neck as described elsewhere²⁶.
7. Attach the tracheal cannula to an end tidal CO₂ monitor and the ventilator delivering 2.5–3.0% isoflurane (depending on the size of the animal) and a 30% O₂ inhalation supplement. Make sure the respiratory rate, inspiratory time, and minute ventilatory volume are set and monitored to ensure an expired end tidal CO₂ of approximately 35 mmHg throughout the experiment.
   NOTE: This is generally achieved with a respiratory rate of approximately 48–60 breaths/min, a tidal volume of 1.70–2.30 mL, and an inspiration time of 0.50–0.60 s for a 250–300 g rat.
8. Fill two PE50 polyethylene cannulas with 1 U/mL heparin in isotonic NaCl solution to prevent clotting and to maintain patency of the catheters. After filling, bevel the open end of each cannula with surgical scissors to facilitate insertion into the artery.
9. Cannulate the right and left femoral arteries as described elsewhere²⁷ to allow continuous monitoring of arterial pressure in one catheter and blood withdrawal from the other catheter.
   1. After carefully separating the arteries from the surrounding tissue under a dissecting microscope, ligate the distal end of the artery and place two additional sutures around the middle and proximal ends of the artery without tightening the knots.
   2. Use the proximal suture as a lifting ligature to prevent bleeding from the artery after the incision for cannula insertion (step 2.11).
10. Insert a V-shaped wire fashioned from a paper clip under the artery in order to occlude the vessel until the cannula is secured.
11. Under a dissecting microscope make a small incision in the femoral artery near the distal ligation using Vannas scissors. Insert the beveled end of the cannula into the incision and advance it into the femoral artery. Tighten the knot on the middle ligature to secure the cannula in place so it is not dislodged by arterial pressure when the lifting ligature or paper clip is removed.
12. After the middle ligature is tightened, release the tension on the lifting ligature and/or remove the paper clip, and tighten the proximal ligature.
13. Close the incision with fine sutures (3–0 silk) or a surgical staple. Alternatively, place a moist gauze over the incision site, depending on the size of the incision.

3. Skull thinning for LDF measurements

1. Immediately after the cannulas are in place, place the animal in a sternal position and secure the head in a stereotaxic device, being careful not to dislodge the catheters or tracheal tube.
2. Use surgical scissors to make an elliptical incision in the skin covering the cranium. Use a cotton swab to remove any connective tissue, ensuring that the cranium is clean and dry. Place a small elongated and rolled piece of tissue paper around the incision on the scalp to stop any bleeding.
3. Under the dissecting microscope, use a Dremel tool or a dental drill with a 2.15 mm drill bit to thin a small area of bone (approximately 0.5–1 cm depending on the size of the rat) in the parietal area over the left or right somatosensory cortex.
   CAUTION: Thin the bone slowly and carefully to avoid penetrating the skull. While performing this step, saline solution should be applied liberally to prevent the area from overheating.
4. Once the skull has been thinned and the area has a pink appearance and/or blood vessels are visualized, cover the area with mineral oil and use a micromanipulator to position the laser Doppler probe over the exposed cerebral microcirculation so that the tip of the probe is just touching the top of the pool of mineral oil (Figure 1).
   NOTE: It is essential to take LDF measurements in an area where there are no external vibrations that would interfere with the laser Doppler readings and that the probe is securely fixed over the same target area throughout the experiment.

4. Assessing cerebral vascular autoregulation

1. Once the LDF probe is fixed in position, allow a 30–45 min equilibration period before beginning the experiment. After the equilibration period, measure the mean arterial pressure (MAP) and laser cerebral blood flow (LCBF) every 30 s for 2 min and average the values to obtain the baseline values for the prehemorrhage blood pressure and LCBF.
2. To evaluate the cerebral vascular autoregulation in response to arterial pressure reduction, measure the LCBF and MAP following successive withdrawals of 1.5 mL of blood from the femoral artery¹¹. To keep the catheter patent, ensure that a volume of heparin solution (100 U/mL in isotonic saline) approximately equal to the catheter volume is infused after each blood draw.
   NOTE: When infusing the heparin solution to maintain catheter patency, it is important to match the volume of the heparin solution to the volume of the catheter as closely as possible to prevent the animal from receiving too much heparin, which could cause unwanted bleeding.
3. After each blood volume withdrawal, allow the rat to equilibrate for 2 min, after which the MAP and LCBF are recorded every 30 s for 2 min. Repeat the blood volume withdrawals until the animal reaches a MAP of approximately 20 mmHg.
4. Determine the effective autoregulatory range by identifying the range of blood pressures from the prehemorrhage MAP to the LLA (steps 4.5 and 5.3, below).
5. Determine the LLA by identifying the lowest pressure at which LCBF still returns to within 20% of the prehemorrhage control value following blood volume withdrawal, as previously described^11,28 or by identifying the intersection point of the regression lines determined during the plateau phase of autoregulation and below the LLA, where LCBF decreases with each successive blood withdrawal (step 5.3, below).
   NOTE: The criteria for defining the LLA and autoregulatory plateau may differ between laboratories (e.g., Takada et al.²⁸ vs. Jones et al.²⁹) as well as procedures for reducing arterial blood pressure (e.g., withdrawal of a specific volume of blood vs. controlled hemorrhage to reach specific arterial pressure levels)¹¹.
6. At the end of the experiment, euthanize the animal by creating a bilateral pneumothorax while under a surgical plane of anesthesia, as approved by the IACUC.
7. LDF values obtained in the tissue after the animal is euthanized will provide the zero baseline flow value for the experimental setup.

5. Statistical analysis

1. Perform linear regression analysis to evaluate the correlation between the LDF values and their corresponding arterial pressure. Use the baseline LDF readings obtained after the animal is euthanized to ensure that there was no nonspecific LDF signal affecting the measured flow rates.
2. Calculate the LLA using the intersection between the regression lines above and below the autoregulatory plateau. To calculate the LLA using this method, combine the two regression equations and solve the resulting equation for arterial pressure.
3. When comparing different experimental groups, use linear regression analysis to calculate the slopes of the LDF vs. arterial pressure relationship above and below the LLA for each animal and summarize them as mean ± SEM for the animals in that experimental group.

Results

Figure 2 summarizes the results of experiments conducted in 10 male Sprague-Dawley rats fed standard laboratory chow. In those experiments, mean LCBF was maintained within 20% of the prehemorrhage value following the first three blood volume withdrawals, until the mean arterial pressure reached the LLA. Subsequent blood volume withdrawals at pressures below the LLA caused a progressive reduction of LCBF, showing that the cerebral circulation was no longer able to produce a sufficient level o...

Discussion

Evaluation of Tissue Blood Flow Responses with Laser Doppler Flowmetry (LDF). As noted above, the LDF signal is proportional to the number and velocity of moving particles, in this case RBC, in the microcirculation. LDF readings in different organs are well correlated with whole organ blood flow assessed by established methods such as electromagnetic flow meters and radioactive microspheres³⁰ and are generally consistent with studies evaluating the regulation of active tone in can...

Materials

Name	Company	Catalog Number	Comments
3-0 braided black silk suture	Midwest Vet	193.73000.2
Arterial Pressure Transducer	Merit Medical	041516504A
Automated Data Acquisition Systems (WINDAQ & BIOPAC system)	DATAQ Instruments
Blood Pressure Display Unit	Stoelting	50115
Circulating warm water pump	Gaymar Industries	T-pump
End-tidal CO2 monitor	Stoelting	Capstar-100
Heparin Sodium	Midwest Vet	191.46720.3
Kimwipe	Fisher Scientific	06-666A
Laser Doppler Flow Meter	Perimed	PeriFlux 5000 LDPM
Laser Doppler Refill Motility Standard	Perimed	PF1001
Polyethylene Tubing (PE240) (for trachea cannula)	VWR	63018-828
Polyethylene Tubing (PE50) (for femoral catheters)	VWR	63019-048
Rodent Ventilator	Cwe/Stoelting	SAR-830/P
Saline	Midwest Vet	193.74504.3
Sprague-Dawley Outbred Rats	Variable	N/A	Rats were ordered from various companies
Standard Rat Chow	Dyets, Inc.	113755
Stereotaxic Instrument	Cwe/Stoelting	Clasic Lab Standard