We sincerely thank Dr. Guanyi Lu from the Division of Vascular Surgery and Endovascular Therapy at the University of Florida for the technical support on the development of the iliac AVF model, as well as surgical training, and Ravi Kumar from the Department of Applied Physiology and Kinesiology at the University of Florida for the technical support getting the live microsurgical images.
This work was supported by grants from the National Institutes of Health and National Heart, Lung, and Blood, Institute numbers R01-HL148697 (to S.T.S.), as well as the American Heart Association grant number POST903198 (to K.K.).

All procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Florida and Malcom Randall Veterans Affairs Medical Center.
NOTE: Young adult (8-10 weeks old) male C57BL/6J mice were purchased from The Jackson Laboratory and housed in a light (12 h light: 12 h dark cycle), temperature (22 &#176;C &#177; 1 &#176;C), and humidity (50% &#177; 10%) controlled animal facility. Five mice were allowed to dwell per cage (W:18 cm x L:29 cm x H:12.5 cm) with nesting materials, food, and water being made available ad libitum. Following 7 days of habitat acclimation with standard chow, the mice were changed to a casein-based chow diet for 7 days as a diet transition phase. Thereafter, mice were fed the casein-based chow with 0.2%-0.15% adenine supplementation for 2-3 weeks to induce renal dysfunction (CKD) prior to the AVF surgery as previously described22,23,24. Control mice received a casein-based chow diet without adenine supplementation (control). The control and CKD diets were maintained throughout the postoperative recovery period (POD).
1. Pre-operative measurements
<ol>
	<li>Assess baseline/pre-operative outcome measurements, aortoiliac vessel diameters and hemodynamic flow parameters using duplex ultrasound imaging and the hindlimb perfusion via laser Doppler as previously described25.</li>
	<li>Determine the unilateral hindlimb grip strength and treadmill gait appraisal to establish baseline hindlimb function as previously described25,26.</li>
	<li>Assess kidney function by measuring glomerular filtration rate (GFR) via FITC-inulin clearance and/or serum blood urea nitrogen (BUN) level as previously described22,24,27.</li>
</ol>
2. Surgical preparation
<ol>
	<li>Prepare the following surgical tools and supplies (Table of Materials): a hot bead sterilizer, eye lubricant, a pen trimmer, alcohol preps, chlorhexidine wipes, extra-fine Graefe forceps, sterilized 0.9% saline, 29 G and 31 G needle syringes, 2 x 2 non-woven sponges, medium single-ended round (SC-9) and small double-ended hard, sharp, pointed (SC-4) cotton swabs, low-temperature cautery, straight Dumont forceps, 45&#176; angled Dumont forceps, straight Vannas spring scissors, curved Vannas spring scissors, round handled needle holders, multiple sizes of sutures (4-0 silk, 5-0 PGA, 6-0 silk, and 10-0 nylon sutures), heparin, absorbable gelatin sponge, a straight needle holder, and buprenorphine. 
	NOTE: Extremity fixing rubber bands and retractors for the abdomen and legs were handmade.</li>
	<li>Sterilize surgical preps using autoclave with steam sterilization at 120-125 &#176;C for 30 min followed by drying for 30 min prior to the surgery. Utilize 70% ethanol cleansing followed by hot bead sterilization (240-270 &#176;C for 3 min) between each animal surgery.</li>
	<li>Prepare sterilized 0.9% normal saline, heparinized saline (100 IU/mL), and buprenorphine (0.01 mg/mL) using 29-31 G needle syringes.</li>
</ol>
3. Anesthesia and positioning
<ol>
	<li>Initiate mouse anesthesia in the induction chamber (0.8 mL/min, 2.5% isoflurane). Once the mouse is adequately anesthetized, place the mouse in a supine position on the surgery station covered by a sterile drape. Taper the isoflurane concentration down to ~1.2% during the shaving and positioning steps.</li>
	<li>Apply the ocular lubricant to protect the eyes from drying during the surgery.</li>
	<li>Using a pen trimmer, shave the abdominal hair for the operation and the leg hair for postoperative perfusion measurements. Clear the hair off the surgical field.</li>
	<li>Fixate the upper and lower extremities with rubber bands and tacks, check the depth of anesthesia by monitoring the toe pinch reflex, and titrate anesthesia as necessary. Perform the respiratory pattern evaluation every 3-5&#160;min throughout the surgical procedure to calibrate the level of anesthesia.</li>
</ol>
4. Exploration of the surgical target area
<ol>
	<li>Clean the shaved skin area several times, alternating between&#160;alcohol prep and&#160;chlorhexidine wipes in a circular pattern to disinfect the surgical field.</li>
	<li>Make a midline laparotomy from the lower edge of the sternal margin to the pubis symphysis. Dissect out the pubis fat pad to obtain a wider operative field.</li>
	<li>Open the celiotomy to access the peritoneal contents with retractors and eviscerate the small and large intestines using medium, single-ended round cotton swabs. Cover the bowels with a saline-soaked non-woven sponge.</li>
	<li>Once adequate exposure of the retroperitoneal vasculature is obtained, cover the remaining bowel, kidneys, and ureters with small saline-soaked non-woven sponges. Evacuate a distended bladder by gently squeezing the bladder dome with medium, single-ended round cotton swabs as needed.</li>
	<li>Carefully dissect out the perivascular fascia and adipose tissue from approximately 1 cm proximal to the aortic bifurcation extending to the level of the left iliac bifurcation using straight Dumont forceps and small double-ended hard, sharp, pointed cotton swabs. 
	NOTE: The left iliac artery and vein are left adherent to one another while isolating the arteriovenous structures en masse. This step&#160;will provide sufficient vessel mobilization to facilitate AVF creation.</li>
	<li>If any small venous branches are encountered originating from or converging with the left common iliac vein, ligate them using low-temperature cautery with or without 6-0 silk suture as needed.</li>
	<li>Pass the tip of the angled forceps under the left common iliac vascular bundle and gently spread multiple times to mobilize the vessels from the underlying retroperitoneal musculature (Figure 1A).</li>
</ol>
5. Creation of a common iliac arteriovenous fistula anastomosis
<ol>
	<li>Place two 4-0 silk sutures around the isolated left common iliac arteriovenous bundle and use them as ligatures (e.g., cross-clamps) to the vascular bundle. Create a single knot with each 4-0 silk tie and apply them sequentially from proximal to&#160;distal.</li>
	<li>Make sure that the silk-tie cross-clamps are placed sufficiently far apart to isolate ~2 mm of vessel length, and the sequential application of the suture ligatures will precipitate left iliac vein engorgement.</li>
	<li>Using the 4-0 silk suture strings as handles, rotate the left iliac arteriovenous vascular bundle clockwise and fine-tune the position to temporarily locate the vein anterior to the artery (Figure 1B).</li>
	<li>Make a longitudinal venotomy (~1 mm) with straight Vannas spring scissors and gently flush out residual blood from the venous lumen with 0.9% saline (Figure 1C). Use caution during this step, as a high-pressure saline flush can cause venous disruption. 
	NOTE: The region of red color that remains in the iliac artery after venous flushing provides a visual window for the following step.</li>
	<li>Place an imbricating 10-0 nylon suture through the posterior wall of the vein. 
	NOTE: This portion of the iliac vein should be in immediate apposition to the anterior wall of the iliac artery, and the walls are naturally adherent. The suture should go through both walls and tie the suture down with a single knot (Figure 1D). Note that a small amount of bleeding, which originates from the stagnant blood in the iliac artery, will appear once the needle passes through both walls. If intra-luminal bleeding continues during this step, the silk suture cross-clamps may be too loose and need to be further tightened.</li>
	<li>Grasp the imbricated suture ends and place them under gentle tension to displace the anterior wall from the posterior wall of the iliac artery. Make a ~1.0 mm x 0.3 mm elliptical incision using curved Vannas spring scissors, removing the adherent walls of both the iliac artery and vein. 
	NOTE: The arteriovenous fistula is thereby created once this common channel is established. It is possible to injure the lateral walls of the vein during this step as the posterior iliac vein/anterior iliac artery wall incision is performed through venotomy exposure. Caution should be taken to avoid this complication as this can considerably reduce the fistula diameter and lead to thrombus development.</li>
	<li>Gently flush out residual blood of the exposed arterial lumen with 0.9% saline and heparinized saline (100 IU/mL)28 (Figure 1E).</li>
	<li>Following the creation of the AVF, repair the initial anterior wall venotomy using two or three 10-0 nylon sutures in an interrupted fashion (Figure 1F).</li>
	<li>Restore the vascular bundle to its original anatomic orientation and place a small piece of saline-soaked absorbable gelatin sponge adjacent to the repaired venotomy to facilitate hemostasis.</li>
	<li>Loosen the 4-0 single-knot cross-clamp ligatures sequentially from distal to proximal. Monitor the venotomy site closely for excessive bleeding while loosening each suture.</li>
	<li>If the repair is not adequately hemostatic, reapply the cross clamps and place another 10-0 nylon suture at the site of bleeding. If hemostasis is assured, remove the sutures and then the absorbable gelatin sponge.</li>
	<li>Gently rub the vascular bundle with small double-ended hard, sharp, pointed cotton swabs, which further facilitate the restoration of blood flow. Confirm technical success of the operation using visualization of pulsatile, bright red oxygenated blood entering the iliac vein and mixing with dark venous blood returning from the hindlimb.</li>
	<li>Inject heparinized saline (0.2 IU/g)15 into the IVC for systemic anticoagulation to improve AVF patency outcomes. 
	NOTE: Although this step occurs after the vascular reconstruction (as opposed to the human analog where heparinization occurs prior to vessel cross-clamping), a reduction in intraoperative bleeding and improved AVF patency were observed when performed at this stage of the procedure. Injection into a site covered by fascia and/or fat is preferable to prevent bleeding from the puncture site.</li>
	<li>Re-inspect the surgical site for hemostasis after the injection of heparinized saline. If there are no bleeding concerns, close the midline fascia and then the skin incision with absorbable 5-0 PGA sutures in a running fashion.</li>
	<li>For sham operations, follow all the key steps of the procedure except for AVF formation. Apply a single knot of the 4-0 silk ligature at the proximal end of the left iliac arteriovenous bundle and match the clamp times to AVF surgeries (e.g., ~20 min, depending on microsurgeon proficiency).</li>
</ol>
6. Postoperative care and measurement
<ol>
	<li>After laparotomy closure, measure the blood perfusion of the bilateral tibialis anterior muscles and ventral paws using laser Doppler imaging. 
	NOTE: Unilateral perfusion deficits will confirm fistula diversion of arterial flow (&#34;steal&#34;).</li>
	<li>Administer 0.1 mg/kg buprenorphine subcutaneously and return the mouse to a prewarmed mouse cage with highly absorbable soft bedding with nest.</li>
	<li>Allow the mouse to recover in the prewarmed mouse cage until the anesthesia wears off, which will be obvious when the mouse is ambulatory and interactive (~2 h). During recovery, give the mouse easy access to a moisturized, soft diet.</li>
	<li>Administer buprenorphine and/or subcutaneous saline hydration every 12 h up to 48 h and perform daily monitoring for 5 days post-operatively. Euthanize animals with a deteriorating condition or excessive tissue necrosis, classified as a modified ischemia score &#8805;229.</li>
	<li>Use serial duplex ultrasound assessments to evaluate fistula patency post-operatively; mice with fistula thrombosis are excluded from subsequent analyses unless the purpose of the experiments is to characterize AVF maturation failure.</li>
	<li>Determine other postoperative outcome measurements such as local hemodynamics, grip strength, and gait performance during the recovery time period. Collect fistula and muscle tissues to assess histomorphology at the end of the experiment during the sacrifice25,27.</li>
</ol>

The authors have nothing to disclose.

The prevalence of hemodialysis patients with ARHD following AVF creation has continued to increase30,31. Indeed, unresolved symptomatic complications4,32,33 such as pain, weakness, paresthesia, and/or reduced range of motion can negatively impact patient wellbeing4,32,33,</...

The establishment and preservation of functional vascular access remain an important primary goal for end-stage renal disease (ESRD) patients receiving renal replacement therapy via hemodialysis1. Repeated hemodialysis treatments are necessary to remove waste products, normalize electrolytes, and maintain fluid balance once kidney function becomes inadequate, and thus are necessary for long-term survival2. Therefore, vascular access represents a &#34;lifeline&#34; for patients with ESRD, and autogenous arteriovenous fistula (AVF) placement remains a preferred dialysis access option among this cohort3. However, approximately 30%-60% of hemodialysis patients experience a spectrum of hand disabilities, clinically defined as access-related hand dysfunction (ARHD). The symptoms of ARHD can range from weakness and discoordination to monoplegia and digital gangrene, which can occur early after AVF creation or develop gradually with fistula maturation. Further, ARHD complicates the ESRD treatment schedule, which is associated with poor quality of life, high risk of cardiovascular disease, and increased mortality2,3,4.
Several animal models have been developed to study vascular remodeling induced by hemodynamic alterations following AVF creation5,6,7,8,9,10,11,12,13,14,15. Large animal models with iliac or femoral AVF16,17,18,19,20 and rodent models using either carotid artery-jugular vein anastomosis or infrarenal aorta-inferior vena cava fistula formation are well established to examine the aforementioned aspects of AVF maturation and patency21. For example, venous hypertension, greater luminal diameter, and increased vein wall thickness are signatures of successful AVF maturation, whereas substantial fibrosis of the media and intimal hyperplasia or thrombus development with no changes in flow often characterize AVF failures6,15. However, large animal models lack the experimental flexibility or transgenic capabilities of murine models, while current rodent models do not readily facilitate the investigation of ARHD due to either the anatomic location and/or lack of associated limb pathology. Indeed, due to a lack of an established preclinical animal model that recapitulates the relevant clinical phenotype, research progress to elucidate the pathobiological mechanisms and develop novel therapeutic strategies has remained stagnant, despite a progressive increase in the number of symptomatic ARHD patients. Therefore, the primary aim of this study is to introduce a unique mouse model of ARHD, providing procedural steps of AVF microsurgery and characterization of AVF-related pathophysiology.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.15% Adenine diet</td>
			<td>ENVIGO</td>
			<td>TD.130899</td>
			<td>20% casein, 0.15% adenine, 0.9% P</td>
		</tr>
		<tr>
			<td>0.2% Adenine diet</td>
			<td>ENVIGO</td>
			<td>TD.130900</td>
			<td>20% casein, 0.2% adenine, 0.9% P</td>
		</tr>
		<tr>
			<td>10-0 Nylon suture</td>
			<td>AD surgical</td>
			<td>XXS-N1005T4</td>
			<td></td>
		</tr>
		<tr>
			<td>29 G needle syringes</td>
			<td>Exel International</td>
			<td>14-841-32</td>
			<td></td>
		</tr>
		<tr>
			<td>31 G needle syringes</td>
			<td>Advocate</td>
			<td>U-100 insulin syringe</td>
			<td></td>
		</tr>
		<tr>
			<td>4-0 silk suture</td>
			<td>AD surgical</td>
			<td>S-S41813</td>
			<td></td>
		</tr>
		<tr>
			<td>45-degree angled dumont forceps</td>
			<td>Fine Science Tools</td>
			<td>11253-25</td>
			<td></td>
		</tr>
		<tr>
			<td>5-0 PGA suture</td>
			<td>AD surgical</td>
			<td>PSGU-518R13</td>
			<td></td>
		</tr>
		<tr>
			<td>6-0 silk suture</td>
			<td>AD surgical</td>
			<td>S-S618R13</td>
			<td></td>
		</tr>
		<tr>
			<td>Absorbable gelatin sponge</td>
			<td>ETHICON</td>
			<td>1975</td>
			<td></td>
		</tr>
		<tr>
			<td>Alcohol preps</td>
			<td>Covidien</td>
			<td>5110-cs4000</td>
			<td>70% isopropyl alcohol</td>
		</tr>
		<tr>
			<td>Buprenorphine</td>
			<td>NA</td>
			<td>NA</td>
			<td>0.01 g/mL</td>
		</tr>
		<tr>
			<td>C57BL6/J mice</td>
			<td>Jaxon Laboratory</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Casein diet</td>
			<td>ENVIGO</td>
			<td>TD.130898</td>
			<td>20% casein, 0.9% P</td>
		</tr>
		<tr>
			<td>Cotton swabs</td>
			<td>CONSTIX</td>
			<td>SC-9</td>
			<td>Medium single-ended round cotton swab</td>
		</tr>
		<tr>
			<td>Cotton swabs</td>
			<td>CONSTIX</td>
			<td>SC-4</td>
			<td>Small double-ended hard, sharp, pointed cotton swab</td>
		</tr>
		<tr>
			<td>Curity non-woven sponges (2x2)</td>
			<td>Covidien</td>
			<td>9022</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved Vannas spring scissors</td>
			<td>Fine Science Tools</td>
			<td>15001-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Doppler ultrasound</td>
			<td>VisualSonics</td>
			<td>Vevo 2100</td>
			<td></td>
		</tr>
		<tr>
			<td>Extra fine graefe forceps</td>
			<td>Fine Science Tools</td>
			<td>11150-10</td>
			<td>2 pairs</td>
		</tr>
		<tr>
			<td>Eye lubricant</td>
			<td>CLCMEDICA</td>
			<td>Optixcare eye lube</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin (5000 U/mL)</td>
			<td>National Drug Codes List</td>
			<td>63739-953-25</td>
			<td>100 IU/mL</td>
		</tr>
		<tr>
			<td>Hot bead sterilizer</td>
			<td>Fine Science Tools</td>
			<td>18000-50</td>
			<td></td>
		</tr>
		<tr>
			<td>Low-temperature cautery</td>
			<td>Bovie</td>
			<td>AA04</td>
			<td></td>
		</tr>
		<tr>
			<td>Pen trimmer</td>
			<td>Wahl</td>
			<td>5640-600</td>
			<td></td>
		</tr>
		<tr>
			<td>Powder-free surgical gloves</td>
			<td>Ansell</td>
			<td>7824PF</td>
			<td></td>
		</tr>
		<tr>
			<td>Round handled needle holders</td>
			<td>Fine Science Tools</td>
			<td>12076-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile towel drape</td>
			<td>Dynarex</td>
			<td>DY440-MI</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterilized 0.9% saline</td>
			<td>National Drug Codes List</td>
			<td>46066-807-25</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight dumont forceps</td>
			<td>Fine Science Tools</td>
			<td>11253-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight needle holder</td>
			<td>Fine Science Tools</td>
			<td>FST 12001-13</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight vannas spring scissors</td>
			<td>Fine Science Tools</td>
			<td>25001-08</td>
			<td></td>
		</tr>
		<tr>
			<td>TrizChLOR4</td>
			<td>National Drug Codes List</td>
			<td>17033-279-50</td>
			<td></td>
		</tr>
	</tbody>
</table>

a murine model of hemodialysis access-related hand dysfunction

Chronic kidney disease is a major public health problem, and the prevalence of end-stage renal disease (ESRD) requiring chronic renal replacement therapies such as hemodialysis continues to increase. Autogenous arteriovenous fistula (AVF) placement remains a primary vascular access option for ESRD patients. Unfortunately, approximately half of the hemodialysis patients experience dialysis access-related hand dysfunction (ARHD), ranging from subtle paresthesia to digital gangrene. Notably, the underlying biologic drivers responsible for ARHD are poorly understood, and no adequate animal model exists to elucidate the mechanisms and/or develop novel therapeutics for the prevention/treatment of ARHD. Herein, we describe a new mouse model in which an AVF is created between the left common iliac artery and vein, thereby facilitating the assessment of limb pathophysiology. The microsurgery includes vessel isolation, longitudinal venotomy, creation of arteriovenous anastomosis, and venous reconstruction. Sham surgeries include all the critical steps except for AVF creation. Iliac AVF placement results in clinically relevant alterations in central hemodynamics, peripheral ischemia, and impairments in hindlimb neuromotor performance. This novel preclinical AVF model provides a useful platform that recapitulates common neuromotor perturbations reported by hemodialysis patients, allowing researchers to investigate the mechanisms of ARHD pathophysiology and test potential therapeutics.

Animals exposed to an adenine diet have reduced glomerular filtration rates (control: 441.3 &#177; 54.2 &#181;L/min vs. CKD: 165.1 &#177; 118.3 &#181;L/min, p &#60; 0.05) and increased serum blood urea nitrogen levels (control: 20.39 &#177; 4.2 &#181;L/min vs. CKD: 38.20 &#177; 10.65 &#181;L/min, p &#60; 0.05) compared to the animals that received casein-based chow, confirming the presence of kidney insufficiency prior to arteriovenous fistula surgery.
Validation of A...

Watch this Scientific Journal Video about A Murine Model of Hemodialysis Access-Related Hand Dysfunction at JoVE.com

A Murine Model of Hemodialysis Access-Related Hand Dysfunction

This protocol details the surgical steps of murine common iliac arteriovenous fistula creation. We developed this model to study hemodialysis access-related limb pathophysiology.

Chronic kidney disease is a major public health problem, and the prevalence of end-stage renal disease (ESRD) requiring chronic renal replacement ...

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1.6K Views.  University of Florida. This protocol details the surgical steps of murine common iliac arteriovenous fistula creation. We developed this model to study hemodialysis access-related limb pathophysiology. 

Video: A Murine Model of Hemodialysis Access-Related Hand Dysfunction

Murine Model of Wound Healing

Wound healing and repair are the most complex biological processes that occur in human life. After injury, multiple biological pathways become activated. Impaired wound healing, which occurs in diabetic patients for example, can lead to severe unfavorable outcomes such as amputation. There is, therefore, an increasing impetus to develop novel agents that promote wound repair. The testing of these has been limited to large animal models such as swine, which are often impractical. Mice represent the ideal preclinical model, as they are economical and amenable to genetic manipulation, which allows for mechanistic investigation. However, wound healing in a mouse is fundamentally different to that of humans as it primarily occurs via contraction. Our murine model overcomes this by incorporating a splint around the wound. By splinting the wound, the repair process is then dependent on epithelialization, cellular proliferation and angiogenesis, which closely mirror the biological processes of human wound healing. Whilst requiring consistency and care, this murine model does not involve complicated surgical techniques and allows for the robust testing of promising agents that may, for example, promote angiogenesis or inhibit inflammation. Furthermore, each mouse acts as its own control as two wounds are prepared, enabling the application of both the test compound and the vehicle control on the same animal. In conclusion, we demonstrate a practical, easy-to-learn, and robust model of wound healing, which is comparable to that of humans.

A murine model of cutaneous wound healing that can be used to assess therapeutic compounds in physiological and pathophysiological settings.

Wound healing and repair are the most complex biological processes that occur in human life. After injury, multiple biological pathways become activated. ...

A New Murine Model of Endovascular Aortic Aneurysm Repair

Endovascular aneurysm exclusion is a validated technique to prevent aneurysm rupture. Long-term results highlight technique limitations and new aspects of Abdominal aortic aneurysm (AAA) pathophysiology. There is no abdominal aortic aneurysm endograft exclusion model cheap and reproducible, which would allow deep investigations of AAA before and after treatment. We hereby describe how to induce, and then to exclude with a covered coronary stentgraft an abdominal aortic aneurysm in a rat. The well known elastase induced AAA model was first reported in 19901 in a rat, then described in mice2. Elastin degradation leads to dilation of the aorta with inflammatory infiltration of the abdominal wall and intra luminal thrombus, matching with human AAA. Endovascular exclusion with small covered stentgraft is then performed, excluding any interactions between circulating blood and the aneurysm thrombus. Appropriate exclusion and stentgraft patency is confirmed before euthanasia by an angiography thought the left carotid artery. Partial control of elastase diffusion makes aneurysm shape different for each animal. It is difficult to create an aneurysm, which will allow an appropriate length of aorta below the aneurysm for an easy stentgraft introduction, and with adequate proximal and distal neck to prevent endoleaks. Lots of failure can result to stentgraft introduction which sometimes lead to aorta tear with pain and troubles to stitch it, and endothelial damage with post op aorta thrombosis. Giving aspirin to rats before stentgraft implantation decreases failure rate without major hemorrhage. Clamping time activates neutrophils, endothelium and platelets, and may interfere with biological analysis.

Histological and biochemical modifications after aneurysm endograft exclusion are still unclear. We describe a new model of endograft implantation on a murine aneurysm. Through thrombus analysis with and without persistant circulating blood stream interface, and new endovascular biomaterials evaluation, this model will help to better understand endovascular aneurysm exclusion pathobiology.

Endovascular aneurysm exclusion is a validated technique to prevent aneurysm rupture. Long-term results highlight technique limitations and new aspects of ...

A Murine Model of Subarachnoid Hemorrhage

In this video publication a standardized mouse model of subarachnoid hemorrhage (SAH) is presented. Bleeding is induced by endovascular Circle of Willis perforation (CWp) and proven by intracranial pressure (ICP) monitoring. Thereby a homogenous blood distribution in subarachnoid spaces surrounding the arterial circulation and cerebellar fissures is achieved. Animal physiology is maintained by intubation, mechanical ventilation, and continuous on-line monitoring of various physiological and cardiovascular parameters: body temperature, systemic blood pressure, heart rate, and hemoglobin saturation. Thereby the cerebral perfusion pressure can be tightly monitored resulting in a less variable volume of extravasated blood. This allows a better standardization of endovascular filament perforation in mice and makes the whole model highly reproducible. Thus it is readily available for pharmacological and pathophysiological studies in wild type and genetically altered mice.

A standardized mouse model of subarachnoid hemorrhage by intraluminal Circle of Willis perforation is described. Vessel perforation and subarachnoid bleeding are monitored by intracranial pressure monitoring. In addition various vital parameters are recorded and controlled to maintain physiologic conditions.

In this video publication a standardized mouse model of subarachnoid  hemorrhage (SAH) is presented. Bleeding is induced by endovascular Circle of  Willis ...

A Preclinical Murine Model of Hepatic Metastases

Numerous murine models have been developed to study human cancers and advance the understanding of cancer treatment and development. Here, a preclinical, murine pancreatic tumor model of hepatic metastases via a hemispleen injection of syngeneic murine pancreatic tumor cells is described. This model mimics many of the clinical conditions in patients with metastatic disease to the liver. Mice consistently develop metastases in the liver allowing for investigation of the metastatic process, experimental therapy testing, and tumor immunology research.

A preclinical, murine model of hepatic metastases performed via a hemispleen injection technique.

Numerous murine models have been developed to study human cancers and advance the understanding of cancer treatment and development. Here, a preclinical, ...

Ischemic Tissue Injury in the Dorsal Skinfold Chamber of the Mouse: A Skin Flap Model to Investigate Acute Persistent Ischemia

Despite profound expertise and advanced surgical techniques, ischemia-induced complications ranging from wound breakdown to extensive tissue necrosis are still occurring, particularly in reconstructive flap surgery. Multiple experimental flap models have been developed to analyze underlying causes and mechanisms and to investigate treatment strategies to prevent ischemic complications. The limiting factor of most models is the lacking possibility to directly and repetitively visualize microvascular architecture and hemodynamics. The goal of the protocol was to present a well-established mouse model affiliating these before mentioned lacking elements. Harder et al. have developed a model of a musculocutaneous flap with a random perfusion pattern that undergoes acute persistent ischemia and results in ~50% necrosis after 10 days if kept untreated. With the aid of intravital epi-fluorescence microscopy, this chamber model allows repetitive visualization of morphology and hemodynamics in different regions of interest over time. Associated processes such as apoptosis, inflammation, microvascular leakage and angiogenesis can be investigated and correlated to immunohistochemical and molecular protein assays. To date, the model has proven feasibility and reproducibility in several published experimental studies investigating the effect of pre-, peri- and postconditioning of ischemically challenged tissue.

The window of the murine dorsal skinfold chamber presented visualizes a zone of acute persistent ischemia of a musculocutaneous flap. Intravital epi-fluorescence microscopy permits for direct and repetitive assessment of the microvasculature and quantification of hemodynamics. Morphologic and hemodynamic results can further be correlated with histological and molecular analyses.

Despite profound expertise and advanced surgical techniques, ischemia-induced complications ranging from wound breakdown to extensive tissue necrosis are ...

A Murine Model of Irreversible and Reversible Unilateral Ureteric Obstruction

Obstruction of the kidney may affect native or transplanted kidneys and results in kidney injury and scarring. Presented here is a model of obstructive nephropathy induced by unilateral ureteric obstruction (UUO), which can either be irreversible (UUO) or reversible (R-UUO). In the irreversible UUO model, the ureter may be obstructed for variable periods of time in order to induce increasingly severe renal inflammation and interstitial fibrotic scarring. In the reversible R-UUO model the ureter is obstructed to induce hydronephrosis, tubular dilation and inflammation. After a suitable period of time the ureteric obstruction is then surgically reversed by anastomosis of the severed previously obstructed ureter to the bladder in order to allow complete decompression of the kidney and restoration of urinary flow to the bladder. The irreversible UUO model has been used to investigate various aspects of renal inflammation and scarring including the pathogenesis of disease and the testing of potential anti-inflammatory or anti-fibrotic therapies. The more challenging model of R-UUO has been used by some investigators and does offer significant research potential as it allows the study of inflammatory and immune processes and tissue remodeling in an injured and scarred kidney following the removal of the injurious stimulus. As a result, the R-UUO model offers investigators the opportunity to explore the resolution of kidney inflammation together with key aspects of tissue repair. These experimental models are of relevance to human disease as patients often present with obstruction of the renal tract that requires decompression and are commonly left with significant residual kidney impairment that has no current treatment options and may lead to eventual end stage kidney failure.

The murine model of irreversible unilateral ureteric obstruction (UUO) is presented together with the model of reversible UUO in which the ureteric obstruction is relieved by anastomosis of the severed ureter into the bladder. These models enable the study of renal inflammation and scarring as well as tissue remodeling.

Obstruction of the kidney may affect native or transplanted kidneys and results in kidney injury and scarring. Presented here is a model of obstructive ...

Pre-clinical Orthotopic Murine Model of Human Prostate Cancer

To study the multifaceted biology of prostate cancer, pre-clinical in vivo models offer a range of options to uncover critical biological information about this disease. The human orthotopic prostate cancer xenograft mouse model provides a useful alternative approach for understanding the specific interactions between genetically and molecularly altered tumor cells, their organ microenvironment, and for evaluation of efficacy of therapeutic regimens. This is a well characterized model designed to study the molecular events of primary tumor development and it recapitulates the early events in the metastatic cascade prior to embolism and entry of tumor cells into the circulation. Thus it allows elucidation of molecular mechanisms underlying the initial phase of metastatic disease. In addition, this model can annotate drug targets of clinical relevance and is a valuable tool to study prostate cancer progression. In this manuscript we describe a detailed procedure to establish a human orthotopic prostate cancer xenograft mouse model.

Prostate cancer is the second most common cause of cancer-related deaths in the United States. An orthotopic cancer model provides a useful approach to understand the biology of prostate cancer and to evaluate the efficacy of therapeutic regimens. This protocol describes detailed steps necessary to establish an orthotopic prostate cancer mouse model.

To study the multifaceted biology of prostate cancer, pre-clinical in vivo models offer a range of options to uncover critical biological information ...

A Mouse Model of Orthopedic Surgery to Study Postoperative Cognitive Dysfunction and Tissue Regeneration

Surgery is commonly used to improve and maintain quality of life. Unfortunately, in vulnerable patients such as the elderly, complications may occur and significantly diminish the outcome. Indeed, after routine orthopedic surgery to repair a fracture, as many as 50% of elderly patients suffer from neurologic complications like delirium. Also, the capacity to heal and regenerate tissue after surgery decreases with age, and can impact the quality of fracture repair and even osseous integration of implants. Thus, a better understanding of mechanisms that drive these age-dependent changes could provide strategic targets to minimize risk for such complications and optimize outcomes. Here, we introduce a clinically relevant mouse model of tibial fracture. The postoperative changes in these mice mimic some of the cognitive impairments commonly observed after routine orthopedic surgery in humans. Briefly, an incision is performed in the right hind limb under strictly aseptic conditions. Muscles are disassociated, and a 0.38-mm stainless steel pin is inserted into the upper crest of the tibia, inside the intramedullary canal. Osteotomy is then performed, and the wound is stapled. We have used this model to investigate the effects of surgical trauma on postoperative neuroinflammation and behavioral changes. By applying this fracture model in combination with parabiosis, a surgical model in which 2 mice are anastomosed, we have studied cells and secreted factors that systemically rejuvenate organ function and tissue regeneration after injury. By following our step-by-step protocol, these models can be reproduced with high fidelity, and can be adapted to interrogate many biologic pathways that are altered by surgical trauma.

This protocol describes a mouse model of orthopedic surgery that has been used to study mechanisms of postoperative neuroinflammation and behavioral changes, and when combined with parabiosis, to study tissue regeneration during aging.

Surgery is commonly used to improve and maintain quality of life. Unfortunately, in vulnerable patients such as the elderly, complications may occur and ...

A Minimally Invasive Model to Analyze Endochondral Fracture Healing in Mice Under Standardized Biomechanical Conditions

Bone healing models are necessary to analyze the complex mechanisms of fracture healing to improve clinical fracture treatment. During the last decade, an increased use of mouse models in orthopedic research was noted, most probably because mouse models offer a large number of genetically-modified strains and special antibodies for the analysis of molecular mechanisms of fracture healing. To control the biomechanical conditions, well-characterized osteosynthesis techniques are mandatory, also in mice. Here, we report on the design and use of a closed bone healing model to stabilize femur fractures in mice. The intramedullary screw, made of medical-grade stainless steel, provides through fracture compression an axial and rotational stability compared to the mostly used simple intramedullary pins, which show a complete lack of axial and rotational stability. The stability achieved by the intramedullary screw allows the analysis of endochondral healing. A large amount of callus tissue, received after stabilization with the screw, offers ideal conditions to harvest tissue for biochemical and molecular analyses. A further advantage of the use of the screw is the fact that the screw can be inserted into the femur with a minimally invasive technique without inducing damage to the soft tissue. In conclusion, the screw is a unique implant that can ideally be used in closed fracture healing models offering standardized biomechanical conditions.

This protocol describes a minimally invasive osteosynthesis technique using an intramedullary screw for standardized stabilization of femur fractures, which can be used to analyze endochondral bone healing in mice.

Bone healing models are necessary to analyze the complex mechanisms of fracture healing to improve clinical fracture treatment. During the last decade, an ...

Isometric Contractility Measurement of the Mouse Mesenteric Artery Using Wire Myography

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and drugs. It is widely used in many studies on the physiological functions of vascular smooth muscle, the pathogenesis of vascular diseases such as hypertension, and the development of smooth muscle relaxant drugs. The mouse is a widely used model animal with a large pool of disease models and genetically modified strains. We introduced this method to measure the isometric contraction of mouse mesenteric artery in detail. A 1.4-mm segment of mouse mesenteric resistance artery was isolated and mounted on a myograph chamber by passing two steel wires through its lumen. After equilibration and normalization steps, the vessel segment was potentiated by a high-K+ solution twice prior to the contraction assay. As an example of the application of this method in drug development, we measured the relaxant effect of a novel natural substance, neoliensinine, isolated from a Chinese herb, embryos of the lotus seed (Nelumbo nucifera Gaertn.) on mouse mesenteric arteries. The vessel segments mounted on the myograph chamber were stimulated with a high-K+ solution. When the force tension reached a stable sustained phase, cumulative doses of neoliensinine were added to the chamber. We found that neoliensinine had a dose-dependent relaxant effect on smooth muscle contraction, thus suggesting that it bears potential activity against hypertension. In addition, as the vessel segment can survive at least 4 hours after mounting and maintain contractility induced by the high-K+ solution for many times, we suggest that the wire myograph system may be used for the time-consuming process of drug screening.

The wire myograph technique is used to investigate vascular smooth muscle functions and screen new drugs. We report a detailed protocol for measuring the isometric contractility of the mouse mesenteric artery and for screening new relaxants of vascular smooth muscle.

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and ...

Methods Article

A Murine Model of Hemodialysis Access-Related Hand Dysfunction