Visualization and Quantification of the Cell-free Layer in Arterioles of the Rat Cremaster Muscle

Summary

This study demonstrates the surgical preparation of the rat cremaster muscle for the visualization of the in vivo cell-free layer. Considerable factors affecting the accuracy of the cell-free layer width measurement are discussed in this study.

Abstract

The cell-free layer is defined as the parietal plasma layer in the microvessel flow, which is devoid of red blood cells. The measurement of the in vivo cell-free layer width and its spatiotemporal variations can provide a comprehensive understanding of hemodynamics in microcirculation. In this study, we used an intravital microscopic system coupled with a high-speed video camera to quantify the cell-free layer widths in arterioles in vivo. The cremaster muscle of Sprague-Dawley rats was surgically exteriorized to visualize the blood flow. A custom-built imaging script was also developed to automate the image processing and analysis of the cell-free layer width. This approach enables the quantification of spatiotemporal variations more consistently than previous manual measurements. The accuracy of the measurement, however, partly depends on the use of a blue filter and the selection of an appropriate thresholding algorithm. Specifically, we evaluated the contrast and quality of images acquired with and without the use of a blue filter. In addition, we compared five different image histogram-based thresholding algorithms (Otsu, minimum, intermode, iterative selection, and fuzzy entropic thresholding) and illustrated the differences in their determination of the cell-free layer width.

Introduction

In vivo animal studies are instrumental to basic science for understanding human physiology and pathology. In particular, in vivo microhemodynamic studies can elucidate the potential impairment of microcirculatory functions altered by abnormal rheological conditions of blood. A number of previous microhemodynamic studies¹ have used the rat cremaster muscle model for visualizing microvascular blood flow. The cremaster muscle is a thin layer of striated muscle surrounding the testes. Thus, the blood flow in the muscle can be visualized with a trans-illumination microscope by means of surgical exposure. This enables us to acquire the in vivo blood flow images without the use of any fluorescence or contrast agents. In addition, the entire blood perfusion of the muscle network can be controlled by reducing the upstream blood flow with abdominal aorta occlusion². Owing to these advantages, the cremaster muscle model has been widely used to investigate the formation of cell-free layer (CFL) in microvessels^1,3.

The CFL width is a prominent hemodynamic parameter in microcirculation, which has been of great interest for its important roles in regulating microcirculatory functions. The CFL is formed by the shear-induced transverse inward migration of red blood cells (RBCs) towards the flow center⁴. Consequently, this migration leads to the depletion of RBCs near the vessel walls, eventually resulting in a cell-free plasma layer. Accordingly, the parietal CFL naturally becomes a diffusion barrier to oxygen (O₂) delivery from the RBC core to the tissues, and to the scavenging of nitric oxide (NO) by the RBCs^5,6. In addition, the production of NO can also be modulated by the dynamic variations of the CFL width^7,8. Therefore, the roles of the CFL in both gas transport and the regulation of homeostasis in microcirculation need to be fully ascertained to better understand blood flow in microcirculation. Recent studies have focused on bridging the hemodynamics and gas transport functions of the CFL in the microcirculation^9-12. Furthermore, a separate set of studies has also investigated how the pathological elevation in RBC aggregation modulates CFL formation and its effect on O₂ and NO bioavailability in tissues^13,14.

The roles of the CFL become more significant in microcirculation where the relative size of the CFL width to the vessel diameter is prominent. This necessitates an effective approach of quantifying the CFL in in vivo blood flow. Particularly, image acquisition and image analysis are the two key components determining the accuracy of CFL width measurement. Successful visualization of tissue blood flow should be preceded by an appropriate surgical preparation of the animal model. Additionally, a proper image analysis technique is needed to overcome the limitations of conventional manual measurements that are mostly induced by human errors^15,16. With advancements in optical instrumentation and computing power for digital image processing, it is now possible to achieve a more accurate and consistent measurement of the CFL width^17-19. Nonetheless, the accuracy of these measurements, being image-based, still ultimately depends on the quality of the images.

Therefore, this study explores the factors influencing the measurement of the in vivo CFL width. We focused particularly on demonstrating the surgical preparation and digital image analysis for measurements of the CFL width in arterioles of the rat cremaster muscle.

Protocol

This study is in accordance with the National University of Singapore Institutional Animal Care and Use Committee (approved protocol no. R15-0225).

1. Surgical Preparation of the Animal Model

1. Vessel cannulations
   1. Anesthetize a male Sprague-Dawley rats (6 - 7 weeks old) weighing (203 ± 20) g with ketamine (37.5 mg/ml) and xylazine (5 mg/ml ) cocktail through intraperitoneal (i.p.) injection (2 ml/kg). Do not recap the needle or remove it from the syringe after the injection.
   2. Once the animal has been anesthetized (confirmed by toe pinching), place it on a heating pad to maintain its body temperature at 37 °C. Gently shave the hair on the scapulae, anterior cervix, lower abdomen, medial hind leg and scrotal sac. Gently restrain the legs using adhesive paper tapes.
   3. Perform all surgical procedures using microdissection scissors and angled forceps while viewing through a stereomicroscope. Place all sharp surgical tools on a puncture-resistant tray to prevent injury during the surgery.
   4. Scrub all surgical sites 3 times with alternating iodine and 70% alcohol before performing the incision. Flush all catheters with 30 IU/ml heparin-saline solution.
   5. Make a 1 - 1.5 cm midline skin incision on the scapulae using a pair of surgical scissors over the right jugular vein. Separate the fascia by blunt dissection to expose the jugular vein and cannulate it with a polyethylene tube (PE-50) filled with heparin-saline using 5-0 silk sutures. Infuse supplemental anesthesia when necessary (1/3 to 1/2 of initial dosage, intravenous (i.v.)) throughout the course of surgery and experiment.
   6. Perform tracheostomy to maintain airway patency. Make a 1 - 1.5 cm incision in the anterior cervical area. Cannulate the trachea using a polyethylene tube (PE-205) with 2-0 silk sutures to secure the catheter in place.
   7. Monitor blood pressure through cannulation at the femoral artery. Make a 1 - 1.5 cm incision at the left medial surface of the hind leg. Separate the femoral artery by blunt dissection. Cannulate the femoral artery with a polyethylene tube (PE-10) filled with heparin-saline using 5-0 silk sutures.
2. Cremaster Muscle Preparation and Flow Visualization
   1. Insert a 5-0 silk suture through the apex of the scrotal sac to extend it. Make an incision along the ventral surface of the scrotal sac. Regularly apply warm isotonic solution (37 °C; pH 7.4) to the exposed muscle.
   2. Remove surrounding connective tissues carefully and thoroughly using a cotton-tipped applicator.
   3. Insert a 5-0 silk suture through the apex of the cremaster muscle. Cut the suture into two pieces of equal length and tie a knot on each side. Cut the muscle between the two knots and stretch it onto a customized transparent Plexiglas platform by gently pulling the suture. Fix the end of the suture onto the platform with blue tack.
      NOTE: Thorough removal of surrounding connective tissues is crucial in obtaining optimum image contrast.
   4. Repeat step 1.2.3 until 5 to 6 fixations are made. Carefully remove the cremaster muscle from the epididymis using high temperature cautery. Superfuse warm isotonic solution to the exposed muscle to prevent dehydration of the tissue.
      1. Surround the cremaster muscle with folded pieces of gauze. Cover the exposed muscle with a polyvinyl film. The gauze pieces with the film form a shallow basin to hold warm isotonic solution for the water-immersion microscope objective (Figure 1A).
   5. Transfer the animal onto the animal stage of an intravital microscope (Figure 1C). Connect the arterial cannulation to a physiological data-acquisition system for continuous pressure monitoring (Figure 1E).
   6. Maintain the muscle temperature at 35 °C with a heating element attached beneath the animal platform (Figure 1B). Place a temperature probe beside the muscle to provide negative feedback to the power controller of the heating element (Figure 1D).
   7. Leave the animal on the stage for 15 min to equilibrate with the environment.
   8. Visualize blood flow under an intravital microscope with a 40X water-immersion objective and a long working condenser.
   9. Choose an unbranched arteriole (< 60 µm) based on a clear image focus and contrast between the RBC core, CFL and vessel walls, in order to focus the microscope on the diametric plane of the blood vessel. Rotate the camera mounted onto the microscope to align the vessel wall vertically.
   10. Record the blood flow using a high-speed video camera at a frame rate of 3,000/sec for 1 sec. Save the recorded video as uncompressed 8-bit grayscale AVI format to preserve the image quality.
       NOTE: A minimum recording frame rate of 3,000 frames/sec is recommended to ensure that the CFL measurement can be performed at least once per RBC under physiological arteriolar flow conditions.
   11. Use a blue filter with peak transmission at a wavelength of 394 nm and spectral bandpass at 310 - 510 nm to enhance the contrast between the RBCs and plasma.
       NOTE: Ensure that the light spectrum passing through the blue filter from the microscopic light source (100 W halogen lamp) is of low light intensity to prevent any potential tissue damage.
   12. At the end of the experiment, euthanize the animal with an overdose of pentobarbital sodium.

2. Image Analysis

1. Preprocessing for the CFL width measurement
   1. Open MATLAB and run the 'CFL_pre.m' file. (This and other MATLAB files can be found in the Supplemental MATLAB Archive.)
   2. Click 'Open file' to select the video file to analyze.
   3. Adjust the 'Rotation' slider to align the vessel walls vertically.
      NOTE: Users can display the assisting grid lines for the vessel alignment by selecting the 'Grid On' radio button, and adjust the zoom level of the image by sliding the 'Zoom' slider.
   4. Click the 'Confirm Editing' button to confirm the vessel alignment.
   5. Click the 'Set ROI to crop' button to define the region of interest (ROI). The aligned image will be displayed in a pop-up window. Adjust the rectangular objective on the image, and double click to confirm the ROI. Skip this step if the cropping of image is not required.
      NOTE: Only include a single vessel in the ROI to analyze the CFL width from the vessel. Click the 'Reset Image' button to restore the image to its original form, if necessary.
   6. Click the 'Extract Images' button to extract all edited video frames into consecutive bit map images (8-bit grayscale 'bmp' format). The extracted images can be found in the folder with the same name as the selected video file.
2. Measurement of CFL width
   1. Open MATLAB and run the 'CFL_measure.m' file.
   2. Click 'Select Folder' to select the folder containing the extracted images.
   3. Click on the folder containing the images and click 'Select Folder'. The first image frame in the folder will be loaded and shown in the 'Grayscale image' panel, along with its gray intensity histogram in the 'Image Histogram' panel.
   4. Select the desired image frame from the list box to perform the analysis, otherwise the first image frame will be selected.
   5. Click 'Find Vessel Walls' to identify the inner vessel wall in the image, which is determined at the location where the light intensity profile peak transits from dark to light over two pixels.
   6. Check 'Median Filter' to apply a median filter to the image to reduce the 'salt and pepper' noise.
   7. Check 'Auto Contrast' to adjust the image intensities digitally to enhance the image contrast.
   8. Select a thresholding algorithm in the list box which automatically determines a thresholding value (τ) that divides the gray levels into two classes — white pixels with gray levels above τ (CFL), and black pixels with gray levels below τ (RBC core).
      NOTE: As an alternative method, use manual thresholding if none of automated-thresholding algorithm provides an appropriate image thresholding. Click on the 'Manual' radio button and adjust the slider to define the manual thresholding value.
   9. To measure the spatial variation of the CFL widths, enter the pixel resolution in the 'Pixel Resolution' box (the resolution with this experimental setup was 0.42 µm/pixel).
   10. Click the 'Calculate' button to obtain the spatial variation of the CFL widths. Click 'Export .csv' to export the CFL width data in a tabulated format.
   11. To measure the temporal variation of the CFL widths at a specific analysis line along the vessel, click on the 'Temporal Variation' radio button and enter the frame rate information (the frame rate used in this experimental setup was 3,000 frames/sec).
   12. Enter the first frame and last frame of the images for the analysis in the 'Start Frame' and 'Last Frame' boxes, respectively.
   13. Select the position of the analysis line along the vessel by sliding the 'Analysis Line' slide bar. Confirm the position of the analysis line, which is illustrated on both the 'Grayscale image' and 'Binary image'.
   14. Click 'Calculate' to obtain the temporal variation of the CFL widths. Click 'Export .csv' to export the CFL width data in a tabulated format.

Results

The visualization of the CFL in vivo is largely dependent on the surgical preparations of the animal. Excessive blood loss or extended surgery duration may subject the animal to shock and blood flow aberrations. Maintenance of tissue temperature using a heating pad as well as a customized platform during the surgery and experiment is also crucial for maintaining the physiological conditions of the rat. By using a 100 W halogen lamp in the microscope system, no discernible tissue damage was observed even at the e...

Discussion

The measurement of CFL width is essential for a better understanding of the hemodynamics in the microcirculation. In particular, the measurement of CFL widths has been performed in mesenteric⁶, spinotrapezius²⁴ and cerebral²⁵ microcirculations. Conventional measurement of in vivo CFL widths was restricted to estimations by manual inspection of the recorded video frames. The manual measurements required the averaging of several successive video frames before visually identifying t...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Intravital microscope	Olympus	BX51WI	Equipment
High speed camera	Photron	1024PCI	Equipment
Blue filter	HOYA	B390	Equipment
Pressure sensor & biopac system	Biopac system	TSD104A, MP100	Equipment
Temperature controller	Shimaden	SR 1	Equipment
Plasma Lyte A	Baxter	NDC:0338-0221	Warm in 37 °C water bath before use
Saline 0.9%	Braun
Heparin (5,000 IU/ml)	LEO
PE-10 polyethylene tube	Becton Dickinson	427400	.024" OD x .011" ID
PE-50 polyethene tube	Becton Dickinson	427411	.038" OD x .023" ID
PE-205 polyethene tube	Becton Dickinson	427446	.082" OD x .062" ID
2-0 non-absorbable silk suture	Deknatel	113-S
5-0 non-absorbable silk suture	Deknatel	106-S
Water circulating heating pad	Gaymar
Water bath	Fisher Scientific	Isotemp 205	Equipment
Sterile Cotton Gauze	Fisher Scientific	22-415-468
Cotton-tipped applicators	Fisher Scientific	23-400-124
Dumont Forceps	Kent Scientific	INS14188	Surgical instrument
Micro Dissecting forceps	Kent Scientific	INS15915	Surgical instrument
Iris forceps 1 x 2 teeth	Kent Scientific	INS15917	Surgical instrument
Vessel cannulation forceps	Kent Scientific	INS500377	Surgical instrument
Micro scissor	Kent Scientific	INS14177	Surgical instrument
Iris scissor	Kent Scientific	INS14225	Surgical instrument
Vessel clip	Kent Scientific	INS14120	Surgical instrument
Gemini cautery system	Braintree Scientific	GEM 5917	Surgical instrument