Name: A Murine Model of Hemodialysis Access-Related Hand Dysfunction (Video) | JoVE Journal
Uploaded: 2022-05-31T14:30:00.000+00:00
Duration: 8 min 39 s
Description: Learn to create a murine common iliac arteriovenous fistula model for studying hemodialysis access-related hand dysfunction. This method enables examination of uremia and neuromuscular perturbations from hemodynamic changes.

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5 minutes

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>

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<li>Hassabi, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Comparing+strength+and+range+of+motion+of+the+upper+limb+with+AV+fistula+access+with+the+contralateral+upper+limb+among+patients+treated+with+hemodialysis%5BTitle%5D)+AND+%22Researcher+Bulletin+of+Medical+Sciences%22%5BJournal%5D)">Comparing strength and range of motion of the upper limb with AV fistula access with the contralateral upper limb among patients treated with hemodialysis.</a> Researcher Bulletin of Medical Sciences. 22 (1), 1(2017).</li>
<li>Capitanini, A., Galligani, C., Lange, S., Cupisti, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Upper+limb+disability+in+hemodialysis+patients%3A+evaluation+of+contributing+factors+aside+from+amyloidosis%5BTitle%5D)+AND+%22Therapeutic+Apheresis+and+Dialysis%22%5BJournal%5D)">Upper limb disability in hemodialysis patients: evaluation of contributing factors aside from amyloidosis.</a> Therapeutic Apheresis and Dialysis. 16 (3), 242-247 (2012).</li>
<li>Altintepe, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Physical+disability%2C+psychological+status%2C+and+health-related+quality+of+life+in+older+hemodialysis+patients+and+age-matched+controls%5BTitle%5D)+AND+%22Hemodialysis+International%22%5BJournal%5D)">Physical disability, psychological status, and health-related quality of life in older hemodialysis patients and age-matched controls.</a> Hemodialysis International. 10 (3), 260-266 (2006).</li>
<li>Castaneda, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Resistance+training+to+reduce+the+malnutrition-inflammation+complex+syndrome+of+chronic+kidney+disease%5BTitle%5D)+AND+%22American+Journal+of+Kidney+Diseases%22%5BJournal%5D)">Resistance training to reduce the malnutrition-inflammation complex syndrome of chronic kidney disease.</a> American Journal of Kidney Diseases. 43 (4), 607-616 (2004).</li>
<li>Hurton, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Upper+extremity+complications+in+patients+with+chronic+renal+failure+receiving+haemodialysis%5BTitle%5D)+AND+%22Journal+of+Renal+Care%22%5BJournal%5D)">Upper extremity complications in patients with chronic renal failure receiving haemodialysis.</a> Journal of Renal Care. 36 (4), 203-211 (2010).</li>
<li>Mazumder, M. K., Giri, A., Kumar, S., Borah, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+highly+reproducible+mice+model+of+chronic+kidney+disease%3A+Evidences+of+behavioural+abnormalities+and+blood-brain+barrier+disruption%5BTitle%5D)+AND+%22Life+Sciences%22%5BJournal%5D)">A highly reproducible mice model of chronic kidney disease: Evidences of behavioural abnormalities and blood-brain barrier disruption.</a> Life Sciences. 161, 27-36 (2016).</li>
<li>Jia, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+novel+model+of+adenine-induced+tubulointerstitial+nephropathy+in+mice%5BTitle%5D)+AND+%22BioMed+Central+Nephrology%22%5BJournal%5D)">A novel model of adenine-induced tubulointerstitial nephropathy in mice.</a> BioMed Central Nephrology. 14, 116(2013).</li>
<li>Kieswich, J. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+novel+model+of+reno-cardiac+syndrome+in+the+C57BL%2F+6+mouse+strain%5BTitle%5D)+AND+%22BioMed+Central+Nephrology%22%5BJournal%5D)">A novel model of reno-cardiac syndrome in the C57BL/ 6 mouse strain.</a> BioMed Central Nephrology. 19 (1), 346(2018).</li>
<li>Abassi, Z., Goltsman, I., Karram, T., Winaver, J., Hoffman, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Aortocaval+fistula+in+rat%3A+a+unique+model+of+volume-overload+congestive+heart+failure+and+cardiac+hypertrophy%5BTitle%5D)+AND+%22Journal+of+Biomedicine+and+Biotechnology%22%5BJournal%5D)">Aortocaval fistula in rat: a unique model of volume-overload congestive heart failure and cardiac hypertrophy.</a> Journal of Biomedicine and Biotechnology. 2011, 729497(2011).</li>
<li>Brower, G. L., Levick, S. P., Janicki, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Inhibition+of+matrix+metalloproteinase+activity+by+ACE+inhibitors+prevents+left+ventricular+remodeling+in+a+rat+model+of+heart+failure%5BTitle%5D)+AND+%22American+Journal+of+Physiology-Heart+and+Circulatory+Physiology%22%5BJournal%5D)">Inhibition of matrix metalloproteinase activity by ACE inhibitors prevents left ventricular remodeling in a rat model of heart failure.</a> American Journal of Physiology-Heart and Circulatory Physiology. 292 (6), 3057-3064 (2007).</li>
<li>Francis, B. N., Abassi, Z., Heyman, S., Winaver, J., Hoffman, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Differential+regulation+of+ET+%28A%29+and+ET+%28B%29+in+the+renal+tissue+of+rats+with+compensated+and+decompensated+heart+failure%5BTitle%5D)+AND+%22Journal+of+Cardiovascular+Pharmacology%22%5BJournal%5D)">Differential regulation of ET (A) and ET (B) in the renal tissue of rats with compensated and decompensated heart failure.</a> Journal of Cardiovascular Pharmacology. 44, 362-365 (2004).</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><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

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 ± 54.2 µL/min vs. CKD: 165.1 ± 118.3 µL/min, p &lt; 0.05) and increased serum blood urea nitrogen levels (control: 20.39 ± 4.2 µL/min vs. CKD: 38.20 ± 10.65 µL/min, p &lt; 0.05) compared to the animals that received casein-based chow, confirming the presence of kidney insufficiency prior to arteriovenous fistula surgery.
Validation of AVF patency 
Although intra-ope...

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 ...

a-murine-model-of-hemodialysis-access-related-hand-dysfunction

Research

JoVE Journal

Medicine

The surgical procedure for induction of unilateral hindlimb ischemia is demonstrated, with confirmation of ischemia by laser Doppler perfusion imaging.

Murine Model of Hindlimb Ischemia 

The goal is to produce an arteriovenous fistula that is simple and reproducible. This method does not use sutures or glue adhesive. Therefore the samples can be used with the least amount of foreign materials for analysis.

Technical Aspects of the Mouse Aortocaval Fistula

Here we present a murine model of arteriovenous fistula (AVF) failure in which a clinically relevant anastomotic configuration is incorporated. This model can be used to study the pathophysiology and to test possible therapeutic interventions.

A Novel Murine Model of Arteriovenous Fistula Failure: The Surgical Procedure in Detail

This manuscript provides a detailed two-step surgical procedure to perform mouse 5/6th partial nephrectomy (PNx) with pole ligation. Four weeks after surgery, in comparison with sham-operated mice, the PNx mice developed impaired renal function, anemia, cardiac hypertrophy, cardiac fibrosis, and decreased heart systolic and diastolic function.

A Mouse 5/6th Nephrectomy Model That Induces Experimental Uremic Cardiomyopathy

This article shows the method for the surgical placement in mice of an intraperitoneal catheter attached to an access port that is positioned at the back of the animal. Moreover, it explains the procedure for a 5/6 nephrectomy to resemble the uremic state of PD patients.

Surgical Techniques for Catheter Placement and 5/6 Nephrectomy in Murine Models of Peritoneal Dialysis

An aortocaval fistula was created by puncturing the murine infra-renal aorta through both walls into the inferior vena cava and was followed by creation of a stenosis in its outflow via partial ligation of the inferior vena cava. This reproducible model can be used to study central venous stenosis.

Murine Model of Central Venous Stenosis using Aortocaval Fistula with an Outflow Stenosis

This article describes modifications of a procedure to implant a peritoneal dialysis catheter in a murine model to avoid major technical issues observed with the conventional techniques.

A Retrograde Implantation Approach for Peritoneal Dialysis Catheter Placement in Mice

This protocol presents the establishment and confirmation of a postnatal right ventricular volume overload (VO) model in mice with abdominal arteriovenous fistula (AVF), which can be applied to investigate how VO contributes to postnatal heart development.

Establishment and Confirmation of a Postnatal Right Ventricular Volume Overload Mouse Model

This article presents an ischemic CKD model using unilateral renal IRI with contralateral nephrectomy performed one day prior to sample collection; this protocol enables serum-based functional assessment, high survival, and reproducible CKD pathology at 28 and 42 days, providing a versatile platform for mechanistic and interventional studies.

Mouse Model of Acute to Chronic Kidney Disease Transition Induced by Renal Ischemia/Reperfusion Injury

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

Murine Model of Wound Healing

Learn to create a murine common iliac arteriovenous fistula model for studying hemodialysis access-related hand dysfunction. This method enables examination of uremia and neuromuscular perturbations from hemodynamic changes.

A Murine Model of Hemodialysis Access-Related Hand Dysfunction

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