The authors thank Peter Bollen, head of the Biomedical Laboratory for lending the equipment needed to record the footage seen through the microscopes.

Animals were housed in the University of Southern Denmark Animal Care Facility as per institutional guidelines. All procedures involving animal subjects have been approved by The Animal Experiments Inspectorate, Ministry of Environment and Food of Denmark.
1. Pre-surgery irradiation
NOTE: Pre-surgery irradiation takes place 7 days before surgery.
<ol>
	<li>Induce anesthesia.
	<ol>
		<li>Place the mouse in an induction box and set the vaporizer to 3% isoflurane with an oxygen flow rate of 0.8&#8722;1.2 L/min to induce inhalation anesthesia. 
		NOTE: Alternatively, injectable anesthetics can be used but for the short duration of the irradiation inducing inhalation anesthesia was sufficient. For obtaining the results presented in this article, 9-week old female C57BL6 mice were used.</li>
		<li>Make sure the mouse is fully anesthetized by tail or paw pinch test.</li>
	</ol>
	</li>
	<li>Position the mouse for irradiation.
	<ol>
		<li>If fully sedated, move the mouse from the induction box and place it under the source of radiation in supine position and gently fixate the hind limbs with tape. 
		NOTE: The mouse will remain sedated for the short duration of the radiation.</li>
		<li>Place a 1.5 mm thick lead pad to ensure that only the area that undergoes surgery (i.e., the circular area with a diameter of 25 mm around the knee) gets irradiated.</li>
	</ol>
	</li>
	<li>Administer a dose of 10 Gy radiation at a dose rate of 5.11 Gy/min (100 kVp, 10 mA). 
	CAUTION: Safety precautions must be taken when working with radiation. During this experiment, all irradiation was performed in a radiation insulated room, and the source of radiation was only turned on when all personnel had left and sealed the room.</li>
	<li>Place the mouse back in its cage.</li>
</ol>
2. Equipment setup
NOTE: Surgery should be performed in a room dedicated to surgical procedures. The operative surface must be sterile.
<ol>
	<li>Thoroughly clean all operative surfaces with 70% ethanol. Wear hairnet and coveralls. Use sterile surgical instruments and sterile gloves.</li>
	<li>Prepare anesthesia.
	<ol>
		<li>Draw up 1 mL of fentanyl (0.315 mg/mL), 1 mL of midazolam (5 mg/mL), and 2 mL of sterile water. Use different syringes and needles for the different components.</li>
		<li>Mix fentanyl and sterile water by slowly emptying the syringes into a sterile glass tube. When mixed, add midazolam to complete the working solution.</li>
	</ol>
	</li>
	<li>Prepare analgesia.
	<ol>
		<li>Draw up 0.2 mL of buprenorphine (0.3 mg/mL) and 2 mL of saline.</li>
		<li>Mix the volumes by slowly emptying the syringes into a sterile glass tube to complete the working solution.</li>
	</ol>
	</li>
	<li>Turn on the microscope and make sure that the lighting is sufficient, and that the microscope is well adjusted for the operator&#39;s eyes. 
	NOTE: All surgical procedures should be performed under an operating microscope. A magnification range from 4x&#8722;25x is sufficient.</li>
</ol>
3. Preparation
<ol>
	<li>Weigh the mouse pre-surgery by placing the mouse in an empty container on a cleared scale.</li>
	<li>Administer anesthetic.
	<ol>
		<li>Draw up 0.1 mL of anesthetic per 10 g of mouse bodyweight. Inject the anesthetic subcutaneously as a bolus injection.</li>
		<li>Let the mouse rest in a cage with plenty bedding and shelter for approximately 10 min until fully sedated. Examine the anesthetic depth by assessing muscle relaxation and perform paw or tail pinch test.</li>
	</ol>
	</li>
	<li>When fully sedated, shave the hind limb chosen for the procedure using electrical clippers. Make sure to wipe of excess hair.</li>
	<li>Turn on the heating device, such as a heating pad and cover it with a surgical cloth.</li>
	<li>Set the flow of oxygen to 0.8 L/min and connect it with a nosecone. Use 100% oxygen. 
	NOTE: The nosecone is only for oxygen delivery and not anesthesia.</li>
	<li>Apply ophthalmic ointment and inject 0.5 mL of saline subcutaneously, preferably in the scruff of the mouse, to prevent hypovolemia during surgery.</li>
	<li>Position the mouse for surgery.
	<ol>
		<li>Place the mouse on the surgical cloth in supine position. Place the nosecone over the snout.</li>
		<li>Fixate the end of the hindlimbs gently with tape to prevent the mouse from shifting during surgery.</li>
		<li>Sterilize the skin using alcohol/chlorhexidine or alcohol/povidone iodine.</li>
	</ol>
	</li>
</ol>
4. Surgery
NOTE: In this example, the left hind limb (when the mouse is viewed in supine position), has been chosen for the procedure.
<ol>
	<li>Make a circular incision.
	<ol>
		<li>Lift the skin with smooth forceps and clip a small opening approximately 5 mm proximal to the popliteal fossa.</li>
		<li>Slide sharp scissors into the opening and clip towards the knee so that the incision ends just above the knee. Make sure not to puncture the underlying vessels by lifting the skin with forceps while clipping.</li>
		<li>Move the mouse to prone position and continue to clip from the knee towards the popliteal fossa until the circumferential incision is complete.</li>
	</ol>
	</li>
	<li>Dissect the skin below the knee.
	<ol>
		<li>Gently blunt dissect the area below the knee to a couple of millimeters above the ankle, by slowly opening and closing the microscissors while lifting the skin with forceps.</li>
		<li>Carefully snip remaining visible adhesions using microscissors. Use sterile saline regularly to keep the tissue moist during the whole procedure.</li>
	</ol>
	</li>
	<li>Dissect the skin at the proximal rim of the circumferential incision so that it can be retracted with an elastic retractor. 
	NOTE: The retractor allows the operator a better view of the proximal lymph vessel and prevents the proximal rim from shifting during surgery.</li>
	<li>While still in prone position, rotate the hindlimb gently and fixate it with tape, so that the ischiatic vein is visible from the most proximal point of the exposed area to the most distal point.</li>
	<li>Inject approximately 0.01 mL of Patent Blue V subcutaneously between the second and third toe using a 0.5 mL syringe with a 30 G needle. Gently press the paw a couple of times to distribute the Patent Blue V. Visualize the lymph vessels and lymph node through the microscope as the Patent Blue V fills the lymph vessels. 
	NOTE: If the blue color of the lymph vessels fades during the procedure, gently massage the paw to promote uptake, rather than inject more Patent Blue V. Excess use of Patent Blue V may lead to leakage and coloring of the tissue surrounding the lymph vessels which may compromise the procedure.</li>
	<li>Locate the important structures: the popliteal lymph node (PLN), the two lymph vessels distal to the lymph node (DLV1 and DLV2), and the one lymph vessel proximal to the lymph node (PLV). 
	NOTE: All the lymph vessels can be found adjacent to the ischiatic vein. The proximal lymph vessel is usually found medial to the vein, the two distal lymph vessels are found medial and lateral to the vein. The abbreviations of the structures are used in the accompanying video.</li>
	<li>Magnify to clearly visualize the PLV and ligate it with a 10-0 nylon suture using micro-needle holder and microforceps. Press the paw a couple of times to ensure that no Patent Blue V passes proximal to the suture. 
	NOTE: Trimming the fat surrounding the lymph vessel may be necessary.</li>
	<li>Repeat step 4.7 to ligate the two distal lymph vessels. Press the paw several times to ensure that no Patent Blue V passes proximal to the ligature. If the lymph vessels lie to close to the ischiatic vein, try dissecting even further distally. 
	NOTE: In this example, it can be seen that one of the lymph vessels bursts due to the ligature hindering the lymph flow. The lymph vessels will often split from the vein further down.</li>
	<li>Remove the popliteal lymph node.
	<ol>
		<li>Locate the popliteal lymph node and clip a small hole with microscissors to access it and remove it with microforceps and microscissors. 
		NOTE: The lymph node has a smooth pearl-like surface in contrast to the surrounding fat tissue.</li>
		<li>To test if the removed tissue is a lymph node, place it in a test tube filled with water. 
		NOTE: If the tissue is comprised of fat, the tissue will float. If the tissue is a lymph node, it will sink to the bottom.</li>
	</ol>
	</li>
	<li>Remove the inguinal fat pad and lymph node.
	<ol>
		<li>Before removing the inguinal fat pad, use a bipolar coagulator to cauterize the vessels running through the fat.</li>
		<li>Resect the inguinal fat pad using microforceps and microscissors. Gently clip the cauterized vessels running through the fat. Then gently resect the fat tissue in the inguinal area. 
		NOTE: The lymph node located in the fat is rarely colored by Patent Blue V and can be hard to differentiate from the fat. Removing the fat pad in one piece is the best way to ensure the lymph node has been removed.</li>
	</ol>
	</li>
	<li>Rinse the leg thoroughly with sterile saline and confirm through the microscope that any small hairs and particles has been thoroughly removed from the surgical area to avoid wound contamination and infection. Make sure there is no active bleeding.</li>
	<li>Suture the skin edges down to the muscle facia with a 6-0 nylon suture using forceps and needle holder, leaving a gap of 2&#8722;3 mm to constrain the superficial lymph flow. 
	NOTE: The accompanying video shows an example of finished sutures.</li>
	<li>Administer analgesia. Draw up 0.1 mL of analgesia per 30 g of mouse bodyweight. Inject the analgesia subcutaneously as a bolus injection.</li>
	<li>Weigh the mouse for post-surgery for comparison.</li>
	<li>Place the mouse in a cage in a cabinet heated for recovery.</li>
</ol>
5. Postoperative care
<ol>
	<li>Give the mice individual cages to recover after surgery with water and food ad libitum.</li>
	<li>Administer a bolus subcutaneous dose of 0.02 mL of buprenorphine 3x daily for 3 days for analgesia.</li>
	<li>Monitor the animal daily for appropriate wound healing, signs of pain and infection. If signs of infection are present, use antibiotic ointment.</li>
</ol>
6. Post-surgery irradiation
<ol>
	<li>Three days after surgery, repeat the procedure for pre-surgery irradiation (steps 1.1&#8722;1.4).</li>
</ol>

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The authors have nothing to disclose.

There are a few critical steps in this protocol. Firstly, it is important that the researchers take safety precautions when working with radioactivity. Secondly, during the surgical part of this protocol, it is important to start the procedure once the mouse has been anesthetized and finish it without unnecessary breaks. This is important to avoid an excessively long surgical period for the animal and to prevent that the anesthesia loses effect during surgery. It is recommended to only administer one bolus injection of a...

Lymphedema is characterized by an accumulation of lymph fluid that leads to localized tissue swelling, which mainly occurs due to impaired or disrupted flow of lymph fluid in the lymphatic vessels1. The lymph flow can be impaired or disrupted by infection, obstruction, injury or congenital defects in the lymphatic system2. These etiologies result in accumulation of lymphatic fluid, which leads to a chronic state of inflammation, resulting in subsequent fibrosis, as well as deposition of adipose tissue3. Lymphedema can be categorized as primary or secondary lymphedema. Primary lymphedema is caused by developmental abnormalities or genetic mutation2,4. Secondary lymphedema occurs due to underlying systemic disease, surgery or trauma2,4. Secondary lymphedema is the most common form of lymphedema in the world2. In developed countries, the most common cause of secondary lymphedema is oncological therapy such as adjuvant radiotherapy and lymph node dissection5. Lymphedema is most frequent among breast cancer patients, but can also develop in patients with gynecologic, melanoma, genitourinary or neck cancer6. It has been suggested that out of all women diagnosed with breast cancer, 21% will develop lymphedema7.
Lymphedema can be stressful to the patient both physically and psychologically. Patients with lymphedema have an increased risk of infection5,8,9, poor quality of life and can develop social anxiety and symptoms of depression10. The complications of chronic lymphedema lead to high cost of care and an increased disease burden9,11. Findings have also suggested that lymphedema might be associated with increased risk of death after breast cancer treatment12. Conservative management such as compression of the affected area, manual lymph drainage and general skincare remain the first line approach. There is currently no curative treatment6. Although progress has been made in the field of surgical and medical therapy, there is still room for improvement. More research, providing insight in the pathophysiology and progression of the disease, is needed to enable clinicians to provide better treatment options for the patients5.
Animal models are being used in preclinical research to understand the pathophysiology of diseases and develop potential treatment options. Several different lymphedema animal models have been established in canines13,14, rabbits15, sheep16, pigs17,18 and rodents19,20,21,22,23,24. The rodent model seems to be the most cost-effective model, when investigating the reconstruction of lymphatic function, due to rodents being easily accessible and relatively low-priced25. The majority of the mice models have focused on inducing lymphedema in the tail of the mice21,22,23. The tail model is very reliable but the exact surgical technique for inducing lymphedema varies significantly in previous published material. This results in fluctuations in duration and robustness of the developed lymphedema presented in known litterature25. Different techniques are also being used for inducing lymphedema in the hindlimb model and they also yield varying results, but the hindlimb model might be easier to understand from a translational perspective. Previous lymphedema models have been hampered by spontaneous lymphedema resolution and therefore a reproducible and permanent lymphedema model is needed25. Researchers have previously tried to increase the dose of radiation, to prevent the spontaneous lymphedema resolution, but this has often led to subsequent severe morbidity25.
This model results in statistically significant lymphedema, without causing severe morbidity, by combining microsurgery with radiation. The model has been revised from a previous surgical model by adding a dose of irradiation that induces lymphedema, without causing severe morbidity26. It also offers a great opportunity for microsurgical training. Having access to microsurgical equipment and a microscope is necessary, due to the small anatomical structures of the mice. The surgical procedure can be performed when the user has been taught basic microsurgical techniques, such as suturing with microsurgical instruments. The operators that performed this procedure all watched tutorial videos by Acland on the preconditions of microsurgical skills (1981) and basic microsuture technique (1985). We recommend practicing the surgical procedure 8&#8722;10 times before using it in research. Practicing the procedure ensures that fewer mistakes are made and that the procedure can be performed more efficiently. When mastered, the surgical procedure can be performed in 45 minutes.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10-0 Nylon suture</td>
			<td>S&#38;T</td>
			<td>12051-10</td>
			<td></td>
		</tr>
		<tr>
			<td>6-0 Nylon suture - Dafilon</td>
			<td>B Braun</td>
			<td>C0933112</td>
			<td></td>
		</tr>
		<tr>
			<td>Coagulator - ICC 50</td>
			<td>ERBE</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton tipped applicators</td>
			<td>Yibon medical co</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting forceps</td>
			<td>Lawton</td>
			<td>09-0190</td>
			<td></td>
		</tr>
		<tr>
			<td>Elastic retractors</td>
			<td>Odense University Hospital</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Electrical clipper</td>
			<td>Aesculap</td>
			<td>GT420</td>
			<td></td>
		</tr>
		<tr>
			<td>Fentanyl 0,315 mg/ml</td>
			<td>Matrix</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Heating pad - PhysioSuite</td>
			<td>Kent Scientific Corp.</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane 1000mg Attane</td>
			<td>Scan Vet</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane vaporizer - PPV</td>
			<td>Penlon</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Micro jewler forceps</td>
			<td>Lawton</td>
			<td>1405-05</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro Needle holder</td>
			<td>Lawton</td>
			<td>25679-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro scissors</td>
			<td>Lawton</td>
			<td>10128-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro tying forceps</td>
			<td>Lawton</td>
			<td>43953-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Microfine U-40 syringe 0,5ml</td>
			<td>BD</td>
			<td>328821</td>
			<td></td>
		</tr>
		<tr>
			<td>Microlance syringe 25g</td>
			<td>BD</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microlance syringe 27g</td>
			<td>BD</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Midazolam 5 mg/ml (hameln)</td>
			<td>Matrix</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Needle holder - Circle wood</td>
			<td>Lawton</td>
			<td>08-0065</td>
			<td></td>
		</tr>
		<tr>
			<td>Non woven swabs</td>
			<td>Selefa</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Opmi pico microscope F170</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Patent blue V - 25 mg/ml</td>
			<td>Guerbet</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors - Joseph</td>
			<td>BD</td>
			<td>RH1630</td>
			<td></td>
		</tr>
		<tr>
			<td>Siemens INVEON multimodality pre-clinical scanner</td>
			<td>Siemens pre-clinical solutions</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Source of radiation - D3100</td>
			<td>Gulmay</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Stata Statistical Software: Release 15</td>
			<td>StataCorp LLC</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Temgesic - 0,2 mg</td>
			<td>Indivior</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Vet eye ointment - viscotears</td>
			<td>Bausch &#38; Lomb</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

a revised method for inducing secondary lymphedema in the hindlimb of mice

Animal models are of paramount importance in the research of lymphedema in order to understand the pathophysiology of the disease but also to explore potential treatment options. This mouse model allows researchers to induce significant lymphedema lasting at least 8 weeks. Lymphedema is induced using a combination of fractioned radiotherapy and surgical ablation of lymphatics. This model requires that the mice get a dose of 10 Gray (Gy) radiation before and after surgery. The surgical part of the model involves ligation of three lymph vessels and extraction of two lymph nodes from the mouse hindlimb. Having access to microsurgical tools and a microscope is essential, due to the small anatomical structures of mice. The advantage of this model is that it results in statistically significant lymphedema, which provides a good basis for evaluating different treatment options. It is also a great and easily available option for microsurgical training. The limitation of this model is that the procedure can be time consuming, especially if not practiced in advance. The model results in objectively quantifiable lymphedema in mice, without causing severe morbidity and has been tested in three separate projects.

This procedure has previously been used in three separate experiments. All the experiments were made by different lead investigators who all are co-authors of this article. In all three experiments, great care was taken to adhere to the same procedure as described in this protocol. In all three experiments, secondary lymphedema was induced in one hindlimb while the other hindlimb served as a control. Volumes of the hindlimbs were the primary outcome in all three experiments. <strong class...

Watch this Scientific Journal Video about A Revised Method for Inducing Secondary Lymphedema in the Hindlimb of Mice at JoVE.com

A Revised Method for Inducing Secondary Lymphedema in the Hindlimb of Mice

This animal model enables researchers to induce statistically significant secondary lymphedema in the hindlimb of mice, lasting at least 8 weeks. The model can be used to study the pathophysiology of lymphedema and to investigate novel treatment options.

Animal models are of paramount importance in the research of lymphedema in order to understand the pathophysiology of the disease but also to explore ...

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Research

JoVE Journal

Medicine

7.9K Views.  University of Southern Denmark. This animal model enables researchers to induce statistically significant secondary lymphedema in the hindlimb of mice, lasting at least 8 weeks. The model can be used to study the pathophysiology of lymphedema and to investigate novel treatment options.

Video: A Revised Method for Inducing Secondary Lymphedema in the Hindlimb of Mice

A Simple Guide Screw Method for Intracranial Xenograft Studies in Mice

The grafting of human tumor cells into the brain of immunosuppressed mice is an established method for the study of brain cancers including glioblastoma (glioma) and medulloblastoma. The widely used stereotactic approach only allows for the injection of a single animal at a time, is labor intensive and requires highly specialized equipment. The guide screw method, initially developed by Lal et al.,1 was developed to eliminate cumbersome stereotactic procedures. We now describe a modified guide screw approach that is rapid and exceptionally safe; both of which are critical ethical considerations. Notably, our procedure now incorporates an infusion pump that allows up to 10 animals to be simultaneously injected with tumor cells.
To demonstrate the utility of this procedure, we established human U87MG glioma cells as intracranial xenografts in mice, which were then treated with AMG102; a fully human antibody directed to HGF/scatter factor currently undergoing clinical evaluation2-5. Systemic injection of AMG102 significantly prolonged the survival of all mice with intracranial U87MG xenografts and resulted in a number of complete cures. 
This study demonstrates that the guide screw method is an inexpensive, highly reproducible approach for establishing intracranial xenografts. Furthermore, it provides a relevant physiological model for validating novel therapeutic strategies for the treatment of brain cancers.

In order to evaluate novel therapeutic paradigms for the treatment of glioma, physiological relevant models are essential. We utilize an implantable guide screw procedure for establishment of intracranial xenograft models that is more rapid and safer than stereotactic approaches.

The grafting of human tumor cells into the brain of immunosuppressed mice is an established method for the study of brain cancers including glioblastoma ...

The Trier Social Stress Test Protocol for Inducing Psychological Stress

This article demonstrates a psychological stress protocol for use in a laboratory setting. Protocols that allow researchers to study the biological pathways of the stress response in health and disease are fundamental to the progress of research in stress and anxiety.1 Although numerous protocols exist for inducing stress response in the laboratory, many neglect to provide a naturalistic context or to incorporate aspects of social and psychological stress. Of psychological stress protocols, meta-analysis suggests that the Trier Social Stress Test (TSST) is the most useful and appropriate standardized protocol for studies of stress hormone reactivity.2 In the original description of the TSST, researchers sought to design and evaluate a procedure capable of inducing a reliable stress response in the majority of healthy volunteers.3 These researchers found elevations in heart rate, blood pressure and several endocrine stress markers in response to the TSST (a psychological stressor) compared to a saline injection (a physical stressor).3 Although the TSST has been modified to meet the needs of various research groups, it generally consists of a waiting period upon arrival, anticipatory speech preparation, speech performance, and verbal arithmetic performance periods, followed by one or more recovery periods. The TSST requires participants to prepare and deliver a speech, and verbally respond to a challenging arithmetic problem in the presence of a socially evaluative audience.3 Social evaluation and uncontrollability have been identified as key components of stress induction by the TSST.4 In use for over a decade, the goal of the TSST is to systematically induce a stress response in order to measure differences in reactivity, anxiety and activation of the hypothalamic-pituitary-adrenal (HPA) or sympathetic-adrenal-medullary (SAM) axis during the task.1 Researchers generally assess changes in self-reported anxiety, physiological measures (e.g. heart rate), and/or neuroendocrine indices (e.g. the stress hormone cortisol) in response to the TSST. Many investigators have adopted salivary sampling for stress markers such as cortisol and alpha-amylase (a marker of autonomic nervous system activation) as an alternative to blood sampling to reduce the confounding stress of blood-collection techniques. In addition to changes experienced by an individual completing the TSST, researchers can compare changes between different treatment groups (e.g. clinical versus healthy control samples) or the effectiveness of stress-reducing interventions.1

This article describes a protocol for inducing psychological stress in participants, which enables researchers to measure psychological, physiological and neuroendocrine responses to stress within single participants or between groups.

This article demonstrates a psychological stress protocol for use in a laboratory setting.  Protocols that allow researchers to study the biological ...

Inducing Myointimal Hyperplasia Versus Atherosclerosis in Mice: An Introduction of Two Valid Models

Various in vivo laboratory rodent models for the induction of artery stenosis have been established to mimic diseases that include arterial plaque formation and stenosis, as observed for example in ischemic heart disease. Two highly reproducible mouse models&#160;&#8211;&#160;both resulting in artery stenosis but each underlying a different pathway of development&#160;&#8211; are introduced here. The models represent the two most common causes of artery stenosis; namely one mouse model for each myointimal hyperplasia, and atherosclerosis are shown. To induce myointimal hyperplasia, a balloon catheter injury of the abdominal aorta is performed. For the development of atherosclerotic plaque, the ApoE -/- mouse model in combination with western fatty diet is used. Different model-adapted options for the measurement and evaluation of the results are named and described in this manuscript. The introduction and comparison of these two models provides information for scientists to choose the appropriate artery stenosis model in accordance to the scientific question asked.

This video shows two models of intimal plaque development in murine arteries and emphasizes the differences in myointimal hyperplasia and atherosclerosis.

Various in vivo laboratory rodent models for the induction of artery stenosis have been established to mimic diseases that include arterial plaque ...

Carotid Artery Infusions for Pharmacokinetic and Pharmacodynamic Analysis of Taxanes in Mice

When proposing the use of a drug, drug combination, or drug delivery into a novel system, one must assess the pharmacokinetics of the drug in the study model. As the use of mouse models are often a vital step in preclinical drug discovery and drug development1-8, it is necessary to design a system to introduce drugs into mice in a uniform, reproducible manner. Ideally, the system should permit the collection of blood samples at regular intervals over a set time course. The ability to measure drug concentrations by mass-spectrometry, has allowed investigators to follow the changes in plasma drug levels over time in individual mice1, 9, 10. In this study, paclitaxel was introduced into transgenic mice as a continuous arterial infusion over three hours, while blood samples were simultaneously taken by retro-orbital bleeds at set time points. Carotid artery infusions are a potential alternative to jugular vein infusions, when factors such as mammary tumors or other obstructions make jugular infusions impractical. Using this technique, paclitaxel concentrations in plasma and tissue achieved similar levels as compared to jugular infusion. In this tutorial, we will demonstrate how to successfully catheterize the carotid artery by preparing an optimized catheter for the individual mouse model, then show how to insert and secure the catheter into the mouse carotid artery, thread the end of the catheter out through the back of the mouse&#8217;s neck, and hook the mouse to a pump to deliver a controlled rate of drug influx. Multiple low volume retro-orbital bleeds allow for analysis of plasma drug concentrations over time.

This method was developed with the goal of delivering a steady drug solution via the carotid artery, to assess the pharmacokinetics of novel drugs in mouse models.

When proposing the use of a drug, drug combination, or drug delivery into a novel system, one must assess the pharmacokinetics of the drug in the study ...

Minimal Invasive Surgical Procedure of Inducing Myocardial Infarction in Mice

Myocardial infarction still remains the main cause of death in western countries, despite considerable progress in the stent development area in the last decades. For clarification of the underlying mechanisms and the development of new therapeutic strategies, the availability of valid animal models are mandatory. Since we need new insights into pathomechanisms of cardiovascular diseases under in vivo conditions to combat myocardial infarction, the validity of the animal model is a crucial aspect. However, protection of animals are highly relevant in this context. Therefore, we establish a minimally invasive and simple model of myocardial infarction in mice, which assures a high reproducibility and survival rate of animals. Thus, this models fulfils the requirements of the 3R principle (Replacement, Refinement and Reduction) for animal experiments and assure the scientific information needed for further developing of therapeutical strategies for cardiovascular diseases.

A highly reproducible model for myocardial infarction in mice with minimal invasive manipulations is described. The model can be easily performed, resulting in a high reproducibility and survival rate. Thus, the described model will reduce the number of required animals as requested by the 3R principle (Replacement, Refinement and Reduction).

Myocardial infarction still remains the main cause of death in western countries, despite considerable progress in the stent development area in the last ...

An Improved Method for Rapid Intubation of the Trachea in Mice

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity is a commonly used model to study both acute lung injury and fibrosis, and multiple methods have been developed for administering bleomycin (and other toxic agents) into the lungs. However, many of these approaches, such as transtracheal instillation, have inherent drawbacks, including the need for strong anesthetics and survival surgery. This paper reports a quick, reproducible method of intratracheal intubation that involves mild inhaled anesthesia, visualization of the trachea, and the use of a surrogate spirometer to confirm exposure. As a proof of concept, 8-12 week old C57BL/6 mice were administered either 2.0 U/kg of bleomycin or an equivalent volume of PBS, and both damage and fibrotic endpoints were measured post-exposure. This procedure allows researchers to treat a large cohort of mice in a relatively short period with little expense and minimal post-procedure care.

This article presents a rapid and simple method for administering bleomycin directly into the mouse trachea via intubation. Key advantages of this method are that it is highly reproducible, easy to master, and does not require specialized equipment or lengthy recovery times.

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity ...

Assessing Therapeutic Angiogenesis in a Murine Model of Hindlimb Ischemia

Critical limb ischemia (CLI) is a serious condition that entails a high risk of lower limb amputation. Despite revascularization being the gold-standard therapy, a considerable number of CLI patients are not suited for either surgical or endovascular revascularization. Angiogenic therapies are emerging as an option for these patients but are currently still under investigation. Before application in humans, those therapies must be tested in animal models and its mechanisms must be clearly understood. An animal model of hindlimb ischemia (HLI) has been developed by the ligation and excision of the distal external iliac and femoral arteries and veins in mice. A comprehensive panel of tests was assembled to assess the effects of ischemia and putative angiogenic therapies at functional, histologic and molecular levels. Laser Doppler was used for the flow measurement and functional assessment of perfusion. Tissue response was evaluated by the analysis of capillary density after staining with the anti-CD31 antibody on histological sections of gastrocnemius muscle and by measurement of collateral vessel density after diaphonization. Expression of angiogenic genes was quantified by RT-PCR targeting selected angiogenic factors exclusively in endothelial cells (ECs) after laser capture microdissection from mice gastrocnemius muscles. These methods were sensitive in identifying differences between ischemic and non-ischemic limbs and between treated and non-treated limbs. This protocol provides a reproducible model of CLI and a framework for testing angiogenic therapies.

Here, a critical hindlimb ischemia experimental model is presented followed by a battery of functional, histologic and molecular tests&#160;to assess the effectiveness of angiogenic therapies.&#8203;

Critical limb ischemia (CLI) is a serious condition that entails a high risk of lower limb amputation. Despite revascularization being the gold-standard ...

A High-Throughput In Situ Method for Estimation of Hepatocyte Nuclear Ploidy in Mice

When the liver is injured, hepatocyte numbers decrease, while cell size, nuclear size and ploidy increase. The expansion of non-parenchymal cells such as cholangiocytes, myofibroblasts, progenitors and inflammatory cells also indicate chronic liver damage, tissue remodeling and disease progression. In this protocol, we describe a simple high-throughput approach for calculating changes in the cellular composition of the liver that are associated with injury, chronic disease and cancer. We show how information extracted from two-dimensional (2D) tissue sections can be used to quantify and calibrate hepatocyte nuclear ploidy within a sample and enable the user to locate specific ploidy subsets within the liver in situ. Our method requires access to fixed/frozen liver material, basic immunocytochemistry reagents and any standard high-content imaging platform. It serves as a powerful alternative to standard flow cytometry techniques, which require disruption of freshly collected tissue, loss of spatial information and potential disaggregation bias.

We present a robust, cost-effective, and flexible method for measuring changes in hepatocyte number and nuclear ploidy within fixed/cryopreserved tissue samples that does not require flow cytometry. Our approach provides a powerful sample-wide signature of liver cytology ideal for tracking the progression of liver injury and disease.

When the liver is injured, hepatocyte numbers decrease, while cell size, nuclear size and ploidy increase. The expansion of non-parenchymal cells such as ...

A Murine Tail Lymphedema Model

Lymphedema is extremity swelling caused by lymphatic dysfunction. The affected limb enlarges because of accumulation of fluid, adipose, and fibrosis. There is no cure for this disease. A mouse tail model that uses a focal full thickness skin excision near the base of the tail, resulting in tail swelling, has been used to study lymphedema. However, this model may result in vascular comprise and consequent tail necrosis and early tail swelling resolution, limiting its clinical translatability. The chronic murine tail lymphedema model induces sustained lymphedema over 15 weeks and a reliable perfusion to the tail. Enhancements of the traditional murine tail lymphedema model include 1) precise full thickness excision and lymphatic clipping using a surgical microscope, 2) confirmation of post-operative arterial and venous perfusion using high resolution laser speckle, and 3) functional assessment using indocyanine green near infrared laser lymphangiography. We also use tissue nanotransfection technology (TNT) for novel non-viral, transcutaneous, focal delivery of genetic cargo to the mouse tail vasculature.

Lymphedema is extremity swelling caused by lymphatic dysfunction. We describe a chronic murine tail model of lymphedema and the novel use of tissue nanotransfection technology (TNT) for genetic cargo delivery to the tail.

Lymphedema is extremity swelling caused by lymphatic dysfunction. The affected limb enlarges because of accumulation of fluid, adipose, and fibrosis. ...

Laser Doppler Perfusion Imaging in the Mouse Hindlimb

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&#8212;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.

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.

Blood flow recovery is a critical outcome measure after experimental hindlimb ischemia or ischemia-reperfusion. Laser Doppler perfusion imaging (LDPI) is ...