Why perfusion mice

Nerve blood flow is important for peripheral nerve function and nerve blood flow deficits have been implicated in peripheral nerve pathologies, e. Metabolic demands of neural tissue are very high and this is reflected in the tissue blood flow. Peripheral nerve blood flow is comparable to blood flow measured in cortical and spinal cord gray matter, and is higher than in white matter of the spinal cord Zochodne, However, unlike the brain, peripheral nervous tissue is not prone to ischemic injury, partly due to its redundant, segmental blood supply and high overall ischemic tolerance.

It takes as long as 1—3 h of ischemia to induce permanent axonal damage in peripheral nerves Zochodne, The measurement of blood supply in peripheral nerves is challenging, particularly at the capillary level. Although chronic hypoxia has been implicated in the onset and progression of DPN, nerve blood supply has thus far mainly been measured in the larger, more proximal peripheral nerves Tuck et al. In the larger nerves, such as the rat sciatic nerve, absolute nerve blood supply in mL blood per g tissue per minute can be measured by radiotracer and hydrogen clearance methods, while relative nerve blood flow can be estimated over time using laser Doppler flow LDF probes.

The aim of this study was to develop an approach to characterize distal peripheral nerve hemodynamics in mice sural nerves located in the hindlimb using in vivo two-photon laser scanning microscopy TPLSM , based on techniques previously used to study perfusion across individual capillaries in the cerebral cortex in rats Kleinfeld et al.

Because peripheral nerves are prone to temperature changes Dines et al. The carcass was then placed in a sealed plastic bag and immersed in ice water for 30 min to solidify the gelatin. The skin on the lower hindlimb was removed and the sural nerve with its over- and under-laying muscles was dissected and embedded in a mold in optimal cutting temperature compound QPath, VWR Inc.

Images were captured and then analyzed using Fiji Schindelin et al. One section was analyzed for each mouse to determine the cross-sectional area of each nerve fiber, and the number of vessels within endoneureal vessels and directly adjacent to epineureal vessels the nerve fiber.

Then, the total number of vessels per nerve fiber area were calculated, including the large and small vessels. Figure 1. Vasculature of murine sural nerve. A Cross section of murine sural nerve and surrounding tissue at the approximate location of in vivo imaging. Vasculature visualized using perfusion with gelatinized India Ink.

Sural nerve highlighted by a dashed black circle. B Murine sural nerve preparation visualized through surgical microscope after preparation of sural nerve window. Sural nerve highlighted by black dashed bracket. C Maximum intensity projection image of a volumetric two-photon in vivo scan of murine sural nerve and its vasculature. Only vasculature is labeled with fluorescent dye, therefore it appears in white.

The nerve is not labeled, therefore it is not visible, but its location is highlighted with a white dashed bracket. Isoflurane was maintained at 1. Following a small incision in the trachea just below the larynx, a 2. An inguinal skin incision was performed in the left hindlimb to cannulate the femoral artery and vein using 0. The venous catheter was used for fluorescent dextran dye injection, and the arterial catheter for continuous blood pressure and heart rate monitoring BP-1 system, WPI Inc.

Mean arterial pressure MAP and heart rate HR were calculated from the continuous blood pressure curves. Following intubation and catheterization, the mouse was placed in a prone position and the right hindlimb was fixed in a custom leg holder. The holder was made from a 3. The outside of the holder Figure 2A was covered with kapton tape and a The exposed ends of the resistance wire were connected to an adjustable linear power supply GPS, GW Instek, Taiwan which was used to warm the fixed hindlimb to a desired temperature.

Figure 2. Experimental setup and line scan measurements. Black arrows show where power source is connected to resistance wire in order to warm up the leg holder. A Custom made leg holder with resistance wire highlighted in red; B Mouse right hindlimb fixed in the leg holder with an imaging window prepared over the sural neve. Red dotted line and red arrowhead indicate the location of the temperature probe inside of the preparation; C Anesthetized mouse in in vivo two-photon microscope ready for imaging.

Red arrowhead and red dotted line show the location of temperature probe inside the preparation; D Sural nerve of the mouse highlighted by yellow dotted lines. Black square indicates imaging location for in vivo two-photon microscopy; E Vasculature of the sural nerve labeled with Texas red dextran. Sural nerve is highlighted by yellow dotted lines, but appears black as it is not labeled; vessels appear in white due to labeling. F Magnification of a microvessel of sural nerve, where line scan has been performed.

Red line shows scan path along the vessel for measurement of RBCv, blue lines show scan path across the vessel for measurement of vessel diameter; G Typical measurements of RBCv, vessel diameter, RBC flux and RBC linear density acquired for 30 s from line scans; H Example of a line scan acquired for 30 s in a sural nerve microvessel.

Black lines are individual red blood cell shadows. Vessel diameter can be estimated from the transversal part of the line scan in blue , RBCv can be estimated from the axial part of the line scan in red by dividing distance traveled by the individual cell by the time of travel.

Following a popliteal incision, the semitendinous and posterior femoral biceps muscles were isolated by blunt dissection and retracted with sutures to expose the sural nerve without causing bleeding Figures 2B,D. If any muscle bleeding occurred, hemostasis was secured by extra fine cauterizing forceps. Dental cement was applied around the edge of the coverslip, fixing it to the leg holder and thus creating a stable sural nerve window Figure 2B. Following these procedures, a 3 cm long, 0. After the completion of surgical preparation, the mouse was positioned under the objective of the two-photon laser microscope Figure 2C.

These temperatures were chosen, as they typically occur in the hindlimb before surgical exposure, after surgical exposure and window preparation, or after positioning for imaging using water immersion objective, respectively. Using this approach, the blood plasma fraction appears bright in the acquired images Figure 2E , while unlabeled RBC appear as dark shadows Figures 2F,H.

Accordingly, at the start of the in vivo imaging protocol, the temperature was set to one of the three temperatures, and nerve perfusion was measured within 3 min after the temperature had stabilized at the desired level. The time needed for limb temperature to stabilize never exceeded 12 min. Accordingly, in each animal, the selected vessels were scanned at all three temperatures when possible.

In some instances, microscopic shifts in tissue position or technical issues related to TPLSM, prevented data acquisition during all three conditions. RBC dynamics were measured as previously described Gutierrez-Jimenez et al. The line scan path was prescribed both along and across the axis of each vessel. Figure 2F shows a typical example of such a line for the line scan. The scan data acquired along the midline of the vessel, indicated by the red line in Figure 2F , was used to estimate RBCv, flux, and LD.

The scan data acquired along the line perpendicular to the vessel, indicated by the blue lines in Figure 2F , was used to estimate vessel diameter, and to provide an additional estimate of RBC flux and LD.

The signal along each line scan was continuously recorded for 30 s at each of the temperatures. Immediately after imaging, the mice were euthanized by sodium pentobarbital overdose Exagon vet. Vessel diameter, RBCv, flux and linear density were estimated from the line scans using software developed in-house for Matlab Rb, Mathworks Inc.

The line scans acquired during the 30 s were stacked to create a two dimensional raw image with time on Y -axis and distance on X -axis Figure 2H. In these images dark, angled streaks appear on the axial portion of the line scan indicated in red in Figures 2F,H. The angle of each streak shows displacement of the individual RBC on both time and distance axes thus resulting in RBCv.

These angles were determined automatically using Radon transform algorithm. Velocity estimates with signal to noise ratio below three were excluded from the analyses Drew et al. Vessel diameter was estimated from the cross sectional part of the line scan indicated in blue in Figures 2F,H as a full width at half maximum, assuming that the cross section represents the full diameter of the vessel.

Pushing fluids through vascular beds of the circulatory system is a core principle of perfusion. This can be done manually using plastic syringes but requires multiple trials to perfect.

Even after years of practicing manual perfusion, maintaining steady flow rates can still be a challenge. It is imperative to avoid air bubbles or to vary the flow velocity References 1. Animal must be unresponsive before continuing. Make a cm lateral incision through the integument and abdominal wall just beneath the rib cage. Carefully separate the liver from the diaphragm.

Make a small incision in the diaphragm using the curved, blunt scissors. The position and pressure of your finger can aid in the ability to cut the diaphragm. Continue the diaphragm incision along the entire length of the rib cage to expose the pleural cavity.

Place curved, blunt scissors along one side of the ribs, carefully displacing the lungs, and make a cut through the rib cage up to the collarbone. Make a similar cut on the contralateral side. Lifting the sternum away, carefully trim any tissue connecting it to the heart. Clamp the tip of the sternum with the hemostat and place the hemostat over the head. When done properly, the thymus lifts away from the heart along with the sternum, providing a clear view of the major vessels.

Make a small incision to the posterior end of the left ventricle using iris scissors. Pass a gauge blunt- or olive-tipped perfusion needle through the cut ventricle into the ascending aorta. The tip should be visible through the wall of the aorta, and should not reach the aortic arch where the brachial and carotid arteries diverge. Use a hemostat to clamp the heart, this secures the needle and prevents leakage.

If desired, the modified hemostat can be used to clamp the aorta around the needle tip these hemostats remain in place until dissection begins, but are omitted from future illustrations for clarity. Finally, make an incision to the animal's right atrium using iris scissors to create as large an outlet as possible without damaging the descending aorta. At this point the animal is ready to be perfused.

Perfusion Open and attach outlet port to needle base taking care not to introduce any air bubbles. Maintain this pressure throughout the buffer infusion period. Start the timer. Adjust the needle angle. The angle of the needle is critical to the achievement of a maximum flow rate note the flow change with angle adjustment.

Switch the buffer valve blue once buffer is almost finished ml. The fluid should be running clear. The clearing of the liver is an indicator of a good perfusion. The liver should be clear at this point. Indicate time for your records. Fixation tremors should be observed within seconds; this should be considered the true time of fixation. The pressure can gradually be increase up to a maximum of mm Hg 2 to maintain a steady flow rate. Close the outlet valve once the fixative is nearly finished.

Indicate ending time for your records. The rat should be stiff at this stage. The used paraformaldehyde must be collected and stored for disposal according to the regulations of your institution. Dissection 4,11 Remove the head using a pair of scissors.

Make a midline incision along the integument from the neck to the nose and expose the skull. Trim off the remaining neck muscle so that the base of the skull is exposed; remove any residual muscle using scissor or rongeurs. Place the sharp end of a pair of iris scissors into the foramen magnum on one side, carefully sliding the scissors along the inner surface of the skull. Next, make a cut extending to the distal edge of the posterior skull surface.

Make an identical cut on the contralateral side. Carefully slide the scissors along the inner surface of the skull as the tip travels from the dorsal distal posterior corner to the distal frontal edge of the skull, lifting up on the blade as you are cutting to prevent damage to the brain. Repeat for opposite side. Using rongeurs peel the dorsal surface of the skull away from the brain.

Trim away the sides of the skull using rongeurs as well. Using a spatula, sever the olfactory bulbs and nervous connections along the ventral surface of the brain. Gently tease the brain away from the head, trimming any dura that still connects the brain to the skull using iris scissors.

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