Laser Doppler Perfusion Imaging in the Mouse Hindlimb

Summary

Here, we present a protocol that demonstrates the technique and necessary controls for Laser Doppler perfusion imaging to measure blood flow in the mouse hindlimb.

Abstract

Blood flow recovery is a critical outcome measure after experimental hindlimb ischemia or ischemia-reperfusion. Laser Doppler perfusion imaging (LDPI) is a common, noninvasive, repeatable method for assessing blood flow recovery. The technique calculates overall blood flow in the sampled tissue from the Doppler shift in frequency caused when a laser hits moving red blood cells. Measurements are expressed in arbitrary perfusion units, so the contralateral non-intervened upon leg is usually used to help control measurements. Measurement depth is in the range of 0.3-1 mm; for hindlimb ischemia, this means that dermal perfusion is assessed. Dermal perfusion is dependent on several factors—most importantly skin temperature and anesthetic agent, which must be carefully controlled to result in reliable readings. Furthermore, hair and skin pigmentation can alter the ability of the laser to either reach or penetrate to the dermis. This article demonstrates the technique of LDPI in the mouse hindlimb.

Introduction

Skin ulceration with inadequate wound healing is a leading cause of amputations in human patients¹. Adequate wound healing requires higher levels of arterial perfusion than are needed to maintain intact skin, which is compromised in patients with peripheral arterial disease^2,3,4. Several other rheumatologic conditions and diabetes can also lead to disturbed and inadequate skin microcirculation to heal wounds^5,6. Many diabetic patients have concomitant peripheral arterial disease, placing them at especially high risk for amputation. Laser Doppler perfusion imaging (LDPI) is used in clinical situations to evaluate the skin microcirculation, as well as in research situations to evaluate blood flow and blood flow recovery after experimental hindlimb ischemia, ischemia-reperfusion, and microsurgical flaps⁷.

The LDPI system projects a low power laser beam that is deflected by a scanning mirror to move over a region of interest. This differs from Laser Doppler flowmetry, which provides a perfusion measurement for the small area of tissue in direct contact with the flowmetry probe⁸. When the laser beam interacts with moving blood in the microvasculature, it undergoes a Doppler frequency shift, which is photodetected by the scanner and converted to arbitrary perfusion units. Because LDPI is a light-based technique, it is limited in terms of depth of penetration to 0.3-1 mm, meaning that for the most part dermal perfusion is assessed⁷. Dermal flow can be altered by skin temperature and the sympathetic nervous system, which may be affected by various anesthetic agents⁹. Measurements from the optical laser are also affected by ambient lighting conditions, skin pigmentation, and can be blocked by overlying fur or hair⁷.

LDPI is the most commonly used research technique to monitor perfusion recovery after ischemia because it is noninvasive, does not require contrast administration, and has quick scan times allowing data collection on multiple animals. This makes it ideal to help determine whether treatments aimed at therapeutic arteriogenesis or angiogenesis are effective in small animal models. Blood flow recovery after hindlimb ischemia as measured by LDPI correlates well with collateral artery development when assessed by other means such as Microfil casting or micro-CT^10,11. The goal of this protocol is to demonstrate the assessment of hindlimb perfusion using LDPI.

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Protocol

Animal experiments were performed according to a protocol approved by the University of Washington Institutional Animal Care and Use Committee.

1. Scanner preparation

1. Adjust the scanner height so that the distance to the scanned subject is approximately 30 cm.
2. Turn on the imager and launch the associated software.
3. Open the Measurement program. If the software is correctly communicating with the scanner, the infrared laser turn on warning will appear.
4. Calibrate the machine with manufacturer provided standards (not shown in the video and will depend on the specific model of machine being used).
5. Adjust the scanner settings to be appropriate for the background material and lighting setup in the room.
   1. Set the gain levels DC FLUX and CONC, per the manufacturer's instructions (not shown in the video).
   2. Set the Background Threshold by pointing the laser beam at the black background material, and press Auto BK Set.

2. Mouse pre-scanning preparation

1. Set up the isoflurane induction chamber with appropriate scavenging of the waste gas.
   NOTE: Placing the induction chamber on a warming pad will help prevent mouse temperature loss during anesthesia induction.
2. Turn on the homeothermic blanket, which is placed in the scanning area underneath a nonreflective surface (in this case a black neoprene fabric). Set the homeothermic blanket to maintain a body temperature of 37 °C.
3. Position the temperature probe for the homeothermic blanket and lubricant so they are ready for insertion.
4. Place the anesthesia mask and scavenging system in the scanning area.
5. Anesthetize the mouse with an isoflurane vaporizer. Set the oxygen rate to 1 L/min of flow and adjust the isoflurane to 4% for anesthesia induction. Turn on the flow to the anesthesia induction chamber, and the mouse breathing rate will slow. Adequate anesthesia is achieved when the mouse loses its righting reflex.
6. Transfer the mouse to an anesthetic mask/nose cone with attached waste gas scavenger and adjust the isoflurane to 1.5%.
   NOTE: This anesthesia level is generally adequate to keep the mouse lying relatively still during scanning, but is not intended to provide surgical levels of anesthesia, so the depth of anesthesia is not checked. Changing the isoflurane level causes changes in heartbeat, respiration, and dermal perfusion, so a consistent percentage should be used throughout any time course experiment and for all experimental subjects. Alternative anesthetic techniques such as IP injection of ketamine xylazine can also be used, but the same anesthetic technique should be used throughout any time course study as different anesthetics affect dermal perfusion differently.
7. (Optional depending on scanning area) If the planned region of interest to be scanned is covered by fur, use a small electric trimmer or depilatory cream to remove the hair from the region of interest.
   NOTE: The depilatory cream should be completely removed, and the mouse skin dried prior to scanning.
8. Place the mouse in the appropriate scanning position on the black nonreflective surface covering the homeothermic blanket, confirming that both hindlimbs remain on the heat source throughout equilibration and scanning (Figure 1).
   NOTE: It is important to maintain both feet on the homeothermic blanket to prevent regional variation in temperature.
9. Insert the lubricated rectal temperature probe associated with the homeothermic blanket.
10. Equilibrate the mouse temperature to desired scanning temperature (37 °C); approximately 5-10 minutes.
11. Select Scanner Setup, which can be accessed from the top menu or from the scanner setup icon. Adjust the scan area by changing the X-Y coordinates to accommodate the region of interest. Scan speed will depend on the scan resolution. Higher resolution will result in longer scan times. For repeat scanning focusing on global perfusion, as opposed to higher resolution focusing on anatomic perfusion, a scan speed of 4 ms/pixel is adequate.
    ​NOTE: Higher resolution and single scan should be considered if the researcher is attempting to directly study the developing collateral circulation (best imaged in the ventral thigh and calf where it is closer to the skin). Repeated scan at lower resolution/speed (e.g., 4 ms/pixel) is adequate when assessing global perfusion to the end organ of the mouse footpad. The software shown in the video automatically loads the previously used template for scanning area, speed, and resolution when restarted, or it can be retrieved from a stored file if different regions of interest are being used for various experiments.
12. If performing repeat scans, select the Repeat and Line Scan tab. The number of scans can be changed (in this case 3 scans) as well as the repeat interval. The minimum time for the repeat interval would be the estimated scanning time, which is shown in the grayed-out area on the right of the box determined by scan area and scan resolution. Adding a few seconds allows the user to pause and potentially reposition the mouse if needed between scans.

3. Scanning

1. Select the Image Scan tab and select the Mark button. The laser will move to outline the scanning area. Adjust the mouse position so that the target to be scanned is within the marked area.
   NOTE: For footpad or footpad and calf scanning, prone positioning with the hindlimbs extended provides a more consistent region of interest than supine positioning. The femoral artery and saphenous artery and collaterals are very close to the ventral surface of the thigh and calf, so supine positioning is preferred if using these regions of interest.
2. Start repeated measurement by selecting the Repeat Scanning icon and press the Play Button to initiate the scan.
3. Confirm the scanning distance in the pop-up window and click OK to start scanning.
4. Monitor the mouse during scanning for mouse movement; if the mouse moves sufficiently that the hindpaws are no longer in the scanning region in the middle of a scan, restart the scan. Small variations in mouse hindpaw position can be accommodated for in the analysis software.
5. Monitor the mouse temperature during the scanning process as it may fluctuate even with the use of the homeothermic blanket. If there is too much variation in the mouse temperature, this may result in significant variation between scans. Generally, a temperature range of 36.8-37.2 °C will result in acceptable data.
6. Save the captured scan under the Save as window with a file name that includes mouse identifier and timepoint for easier data analysis. Enter mouse and timepoint details if desired in the subject details window.
7. Turn off the isoflurane and remove the rectal temperature probe.
8. Disinfect the rectal temperature probe with 70% ethanol so it is ready for use in the next mouse.
9. Allow the mouse to recover from anesthesia to the point where it displays a righting reflex by flipping from the supine position to the prone position prior to returning it to the cage.
   ​NOTE: Recovery can be carried out either on a warming blanket for isoflurane since recovery is very quick or in a warmed recovery cage for ketamine/xylazine.

4. Capturing LDPI data (Figure 3)

1. Open the imaging review software program.
2. Go to the file menu, open, and locate the saved file.
3. Select the ROI icon from the toolbar.
4. Select the Add Polygon button.
5. Trace the region of interest (ROI) for the control hindlimb using the mouse. Polygon tracing does not have to be exact as the gray background will not be included in the calculated averages.
6. Repeat steps 4.3-4.5 for the surgical hindlimb.
7. Choose the Statistics icon to open the Image ROIs Statistics Results (PU) window.
8. Export the results for Polygon 1 (control hindlimb) and Polygon 2 (surgical hindlimb) to a data collection worksheet via copy/paste.

5. Analysis

1. Capture the data as Surgical/Control ratio for each scan.
2. Use the averaged surgical/control for all three scans for the data point for that particular mouse at that timepoint. Because of biologic variability in the response to hindlimb ischemia, in general 8-10 mice are required per timepoint to achieve reproducible results with ~10% standard error.
   NOTE: Before allowing the mouse to recover from anesthesia, it is worthwhile to perform a quick analysis of the repeated scans to check if the data is too variable (e.g., more than 100-150 perfusion units different between scans 1-3). High variation between repeated scans suggests the mouse was not completely equilibrated during the scan (Figure 2), and a repeat scan can be performed without losing a datapoint, which would occur if the images are not analyzed until a later date. Changing the color palette to optimize the dynamic range of displayed flux values may be necessary to better display scan variation (Figure 2).

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Results

Successful LDPI should result in consistent repeated measures scans, with no more than 100-150 perfusion unit variation (corresponding to about 10% of the usual mean perfusion for the mouse footpad) between the three scans (Figure 2). As demonstrated in Figure 2, repeat scans help determine that the mouse has been appropriately equilibrated so that the ischemic/control ratio best reflects the underlying blood flow as opposed to variation in dermal perfusion caus...

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Discussion

Consistent technique is critical for obtaining reliable results with LDPI. The same anesthetic, temperature settings, mouse position, and region of interest should be used throughout the entire time course. Different anesthetic agents will result in higher or lower perfusion values⁹. Isoflurane anesthesia is convenient because of its rapid onset and emergence as well as overall safety. A consistent percentage of isoflurane should be used as depth of anesthesia with this vasodilatory agent may alte...

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Materials

Name	Company	Catalog Number	Comments
Black nonreflective material			Fabric store, black neoprene recommended by company
F/air cannister	A.M. Bickford Inc	80120
Homeothermic blanket with rigid metal probe	Harvard Apparatus		Also comes with flexible probe, but this is less durable
Isoflurane Anesthesia machine	Drager		Multiple manufacturers
Isoflurane induction chamber	VetEquip	941444	2 L chamber
Moor laser Doppler perfusion imager	Moor Instruments	MoorLDI2-IR	Higher resolution imager (MoorLDI2-HIR)
Mouse Anesthesia nose cone			Multiple manufacturers
Nair	Nair
Oxygen tank			Multiple manufacturers
Surgilube	Multiple distributors