We would like to thank Raymond H. Thornton for his insightful and comprehensive early work on this technique.This study was funded by grant support from the National Cancer Institute (NCI 1R37CA250661-01A1), the Children&#39;s Leukemia Research Association, the Hackensack Meridian School of Medicine, and the HUMC Foundation/Tackle Kids Cancer.

All procedures were performed in accordance with animal care guidelines at the Center for Discovery and Innovation (IACUC protocol 290). For the present study, C57BL/6 mice (female, 4-6 weeks old), C57BL/6 mice (female, 6 months old), J:NU female mice, NOD scid gamma (NSG) female mice, and B6;CAG-luc, -GFP mice were used as the young mouse model, aged mouse model, athymic nude model, immunodeficient model, and bioluminescence cell source, respectively. The mice were obtained from a commercial source (see Table of Materials). This procedure will typically require two people (one to remain sterile while performing the injections and another to handle the mice).
1. Animal preparation
<ol>
	<li>Induce anesthesia in the mice using 3%-4% isoflurane gas and maintain anesthesia using 1%-3% isoflurane gas administered&#160;via&#160;a nose cone and precision-calibrated vaporizer (see&#160;Table of Materials).</li>
	<li>Confirm the appropriate anesthetic depth/unconsciousness by unresponsiveness to the hind paw pinch.</li>
	<li>Remove the fur from the anterior chest area of the mice by applying a thin layer of depilatory cream for less than 1 min. Use a wet paper towel to remove the cream completely along with the loose fur. 
	NOTE: Applying too much cream will result in the chest area skin becoming inflamed.</li>
	<li>Place one mouse at a time, supine, on the heated platform of the small animal ultrasound imaging station (see Table of Materials) with the nose cone in place (Figure 1).</li>
	<li>Secure the mouse to the stage with medical adhesive tape at the hind and forelimbs (Figure 1).</li>
	<li>Apply ophthalmic ointment to both eyes to prevent corneal drying.</li>
	<li>Disinfect the fur-free upper thorax skin using a chlorhexidine gluconate applicator (see Table of Materials).</li>
</ol>
2. Preparation of the ultrasound machine and sterile field
<ol>
	<li>Activate the highest frequency linear probe available, typically the probe with the highest spatial resolution for the size of the animal being imaged. Activate the probe by tapping the corresponding button after the startup screen. 
	NOTE: For this application with mice, the probe used is designed specifically for use with mice and small rats (see Table of Materials).</li>
	<li>Optimize the ultrasound settings for imaging and injection following the steps below.
	<ol>
		<li>Adjust the depth of the field of view to an appropriate size for the target animal by adjusting the vertically oriented sliders on the right side of the screen (Figure 2). The maximum depth setting will typically be about 6-8 mm for young mice.</li>
		<li>Adjust the grayscale gain by sliding the button along the horizontal bar on the bottom of the screen (Figure 2). The goal is to start with an image that is just slightly darker than a typical &#34;gray&#34; appearance.</li>
		<li>Adjust the focal zone (blue arrow on the right of the screen, Figure 2) to the anticipated level of the thymus. For young mice, this will be around a depth of 4 mm.</li>
		<li>If image capture is desired, test the functionality of the store image and store clip buttons to ensure the images can be appropriately saved throughout the procedure. Perform this by tapping the Save Clip button on the bottom right of the screen or by tapping the Freeze button, and then tapping Save Image (Figure 2).</li>
	</ol>
	</li>
	<li>Apply a small amount (~1 mL) of ultrasound gel to the transducer surface (see Table of Materials) while it is upright, either resting in the ultrasound machine holder or in the hands of an assistant.</li>
	<li>Prepare a small sterile field next to the heated platform. The optimal positioning for this is usually between the platform and the ultrasound machine.
	<ol>
		<li>Empty these items onto the sterile field: a sterile probe cover, rubber band, sterile gloves, and sterile ultrasound gel (see Table of Materials).</li>
		<li>With the sterile field set up and items in place, put on the sterile gloves.</li>
		<li>Carefully place the sterile probe cover over the ultrasound transducer (as well as over the gel that was initially placed on the probe). Maintain sterility and touch only the sterile cover, nothing else. Slide the sterile rubber band over the sterile probe cover to keep it in place. 
		NOTE: The air foci, regardless of size, can interfere with ultrasound imaging. Hence, it is essential to apply the ultrasound gel between the transducer and sterile probe cover, and on top of the probe cover to ensure an air-free interface between the ultrasound probe and the animal.&#160;</li>
		<li>Place a moderate amount (2-3 mL) of sterile ultrasound gel onto the transducer. 
		&#8203;NOTE: The user is now ready to image an anesthetized mouse.</li>
	</ol>
	</li>
</ol>
3. Imaging and locating the thymus
<ol>
	<li>While maintaining sterility, place the ultrasound gel-topped probe vertically on the disinfected portion of the anterior chest wall of the mouse for initial imaging.
	<ol>
		<li>Take a moment to look at the ultrasound image and optimize it further. Go back to step 2.2 and adjust to get an appearance similar to that of Figure 3.</li>
	</ol>
	</li>
	<li>Scan the anterior chest of the mouse in a transverse plane. Perform this by holding the transducer vertically and moving it up and down from the neck to the abdomen in a paintbrush-like or &#34;sweeping&#34; motion. 
	NOTE: The heart will be the most recognizable structure in the chest due to its rapid motion and &#34;chambered&#34; appearance. Once the heart is localized, this can be used as a reference point to acquire an image of the thymus.</li>
	<li>With the heart centered in the field of view, sweep the transducer slightly toward the neck. Just superior to the heart, the thymus is usually encountered.</li>
	<li>Visualize the thymus as a bilobed, pyramidal, hypoechoic (&#34;dark&#34; or &#34;black&#34; appearing on the screen) structure that is centered in the midline, anterior to the aorta and posterior to the sternum (Figure 3A).</li>
	<li>Make a note of the two round paired black (i.e., &#34;hypoechoic&#34;) structures on either side of the upper chest. 
	&#8203;NOTE: These are the bilateral venae cavae. The aorta is a similar curvilinear hypoechoic structure in the midline between the two venae cavae. These are easily recognizable by their pulsatile motion.</li>
</ol>
4. Injection of the thymus
<ol>
	<li>If needed, apply more (2-3 mL) sterile ultrasound gel to the transducer. 
	NOTE: A relatively large amount of sterile gel on the transducer (compared to the size of the mouse thorax) will act as a &#34;gel pad&#34; around the chest wall of the mouse. This will reduce the number of ultrasound artifacts made by air within the field of view.</li>
	<li>Using the ultrasound probe, locate the widest portion of the thymus, which is usually the ideal target site for injection. Anticipate a horizontal needle trajectory in the chosen location.
	<ol>
		<li>Note where the major blood vessels (SVCs and aorta) are located at this site. Avoid these during the injection.</li>
		<li>The blood vessels will be hypoechoic, pulsatile structures, as described in step 3.7. If unsure, use the color Doppler mode to check for flow within the vessels (Figure 4A). Activate the color Doppler mode by tapping the Color button on the screen.</li>
		<li>If one of the major blood vessels (or the heart) is anticipated to be along the expected needle trajectory, choose a new target area or find a different approach/trajectory.</li>
	</ol>
	</li>
	<li>Hold the transducer in one hand and a 30 G insulin needle (see Table of Materials) with 10 &#181;L of injectate in the other. 
	NOTE: The injectate will vary based on the experimental design. The present study used phosphate-buffered saline, trypan blue, or D-luciferin (0.1 &#181;g/10 &#181;L).</li>
	<li>To begin the injection process, move the transducer laterally so that the thymus is off-center in the ultrasound field of view. Ensure that the other side of the field of view consists of mostly ultrasound gel and nothing else.</li>
	<li>Place the tip of the needle in the gel under the transducer and slowly move the needle until it is visualized adjacent to the skin surface (Figure 4B).</li>
	<li>While continuously imaging the needle under ultrasound, insert the needle into the thymus gland with a percutaneous trajectory, away from blood vessels.
	<ol>
		<li>Use a &#34;cross thymus&#34; horizontal trajectory to place the needle tip in the thymic lobe contralateral to the entry site. This accounts for potential leakage along the needle tract (Figure 5A).</li>
	</ol>
	</li>
	<li>Once the needle tip is inside the desired portion of the thymus, quickly inject the contents (such as 10 &#181;L of trypan blue or D-luciferin, 0.1 &#181;g/10 &#181;L) from the 30 G syringe while using sonographic visualization.
	<ol>
		<li>To stabilize the syringe during needle insertion and injection, hold the syringe between the thumb and third finger and control the syringe plunger with the index finger.</li>
	</ol>
	</li>
	<li>Remove the needle after all the contents have been deposited.</li>
</ol>
5. Post-injection monitoring of animals
<ol>
	<li>Transfer the animal to an empty cage and observe until it regains sufficient consciousness to maintain sternal recumbency. 
	NOTE: Full recovery from anesthesia is expected to occur within 2 min.</li>
	<li>Monitor the animal for an additional 10 min for signs of distress, labored breathing, or bleeding. 
	NOTE: Post-injection pain is not expected, and there is typically no need for post-injection analgesia.</li>
	<li>Once fully recovered and following an uneventful post-injection observation period, return the injected animal to the company of other animals.</li>
</ol>

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A., Bhandoola, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Circulating+hematopoietic+progenitors+with+T+lineage+potential.">Circulating hematopoietic progenitors with T lineage potential.</a> Nature Immunology. 5 (9), 953-960 (2004).</li><li>Marodon, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Induction+of+antigen-specific+tolerance+by+intrathymic+injection+of+lentiviral+vectors.">Induction of antigen-specific tolerance by intrathymic injection of lentiviral vectors.</a> Blood. 108 (9), 2972-2978 (2006).</li><li>Adjali, O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+correction+of+ZAP-70+immunodeficiency+by+intrathymic+gene+transfer.">In vivo correction of ZAP-70 immunodeficiency by intrathymic gene transfer.</a> Journal of Clinical Investigation. 115 (8), 2287-2295 (2005).</li><li>Tuckett, A. Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Image-guided+intrathymic+injection+of+multipotent+stem+cells+supports+life-long+T+cell+immunity+and+facilitates+targeted+immunotherapy.">Image-guided intrathymic injection of multipotent stem cells supports life-long T cell immunity and facilitates targeted immunotherapy.</a> Blood. 123 (18), 2797-2805 (2014).</li><li>Tuckett, A. Z., Thornton, R. H., O'Reilly, R. J., vanden Brink, M. R. M., Zakrzewski, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intrathymic+injection+of+hematopoietic+progenitor+cells+establishes+functional+T+cell+development+in+a+mouse+model+of+severe+combined+immunodeficiency.">Intrathymic injection of hematopoietic progenitor cells establishes functional T cell development in a mouse model of severe combined immunodeficiency.</a> Journal of Hematology &amp; Oncology. 10 (1), 109 (2017).</li><li>Hogan, B. V., Peter, M. B., Shenoy, H. G., Horgan, K., Hughes, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Surgery+induced+immunosuppression.">Surgery induced immunosuppression.</a> Surgeon. 9 (1), 38-43 (2011).</li><li>Blair-Handon, R., Mueller, K., Hoogstraten-Miller, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+alternative+method+for+intrathymic+injections+in+mice.">An alternative method for intrathymic injections in mice.</a> Laboratory Animals. 39 (8), 248-252 (2010).</li><li>Tuckett, A. Z., Zakrzewski, J. L., Li, D., vanden Brink, M. R., Thornton, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Free-hand+ultrasound+guidance+permits+safe+and+efficient+minimally+invasive+intrathymic+injections+in+both+young+and+aged+mice.">Free-hand ultrasound guidance permits safe and efficient minimally invasive intrathymic injections in both young and aged mice.</a> Ultrasound in Medicine and Biology. 41 (4), 1105-1111 (2015).</li><li>K&#252;ker, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+value+of+necropsy+reports+for+animal+health+surveillance.">The value of necropsy reports for animal health surveillance.</a> BMC Veterinary Research. 14 (1), 191 (2018).</li><li>Sinclair, C., Bains, I., Yates, A. J., Seddon, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Asymmetric+thymocyte+death+underlies+the+CD4:CD8+T-cell+ratio+in+the+adaptive+immune+system.">Asymmetric thymocyte death underlies the CD4:CD8 T-cell ratio in the adaptive immune system.</a> Proceedings of the National Academy of Sciences of the United States of America. 110 (31), 2905-2914 (2013).</li><li>Manna, S., Bhandoola, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intrathymic+injection.">Intrathymic injection.</a> Methods in Molecular Biology. 1323, 203-209 (2016).</li><li>de la Cueva, T., Naranjo, A., de la Cueva, E., Rubio, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Refinement+of+intrathymic+injection+in+mice.">Refinement of intrathymic injection in mice.</a> Laboratory Animals. 36 (5), 27-32 (2007).</li></ol>

The authors do not have any conflicts of interest to disclose.

An ultrasound-guided free-hand injection is a highly accurate technique for delivering study materials to the thymus in an efficient and aseptic fashion. Following the initial sterilization of the skin at the injection site, sterility is maintained during the procedure owing to the use of sterile gloves, sterile ultrasound probe covers, and sterile ultrasound gel. In contrast to the blind percutaneous approach10,17 or relying on surgical incisions for direct visu...

The thymus has an essential role in T cell development and immunity. T cell deficiency, which can be caused by thymic involution, genetic disorders, infections, and cancer treatments, amongst other factors, leads to high mortality and morbidity1,2. Mouse models are indispensable in both basic and translational immunology research and have been used for decades to study thymic biology and T cell development, as well as to develop treatments for those suffering from thymic dysfunction and T cell deficiency3,4,5.
A central part of thymic investigations has been the intrathymic injection of biological materials such as cells, genes, or proteins in mouse models6,7,8,9,10,11,12. Conventional intrathymic injection methods use thoracotomy followed by intrathymic injection under direct visualization or by &#34;blind&#34; percutaneous injection into the mediastinum. The surgical approach significantly increases the pneumothorax risk, amongst others. Moreover, the elevated stress during this surgery results in immunosuppression, thus potentially compromising immunological data13. Experienced researchers, after some practice, can perform the blind injection technique, but this approach is less accurate and therefore, limits experimental subjects to young mice with a big thymus.
The utilization of ultrasound guidance has been introduced as a precise and minimally invasive alternative to traditional intrathymic injection approaches14. However, this procedure is very time-consuming when using the integrated rail system instead of the free-hand technique. Performing injections with the injection mount requires careful imaging optimization and positioning of the transducer with the help of the various attachments such as the transducer stand and mount, the X, Y, and Z positioning system, as well as proficient operation of the micro-manipulation controls and rail system extensions. A simple alternative technique, ultrasound-guided thymic injection, is presented here performed by a radiologist using a free-hand approach15, which is both a rapid and accurate minimally invasive alternative to the above-described methods. Importantly, the current approach can be performed with any high-resolution ultrasound imaging system without needing an injection mount and integrated rail system. It is especially useful for studies requiring the injection of large numbers of mice11, for experiments involving the injection of both thymic lobes, or for the accurate injection of small thymuses in aged, irradiated, or immunocompromised mice12.

<table><tbody><tr><td>Aquasonic 100 Ultrasound Gel</td><td>Parker Laboratories (Fairfield, NJ, USA)</td><td>01-01</td><td>Sterile Ultrasound Transmission Gel</td></tr><tr><td>B6;CAG-luc, -GFP mouse</td><td>The Jackson Laboratory (Bar Harbor, ME, USA)</td><td>025854</td><td>Bioluminescence cell source</td></tr><tr><td>BD Insulin Syringes with needle</td><td>Becton Dickinson (Franklin Lakes, NJ, USA)</td><td>328431</td><td>Ultra-fine needle - 12.7 mm, 30 G</td></tr><tr><td>C57BL/6 mouse - aged</td><td>The Jackson Laboratory (Bar Harbor, ME, USA)</td><td>000664</td><td>age 6 months old; aged model</td></tr><tr><td>C57BL/6 mouse - young</td><td>The Jackson Laboratory (Bar Harbor, ME, USA)</td><td>000664</td><td>age 4-6 weeks; young model</td></tr><tr><td>Chloraprep One-step 0.67 mL</td><td>CareFusion (El Paso, TX, USA)</td><td>260449</td><td>chlorhexidine gluconate applicator</td></tr><tr><td>Curity Cotton Tipped Applicator</td><td>Cardinal Health (Dublin, OH, USA)</td><td>A5000-2</td><td>Sterile, 6&quot;</td></tr><tr><td>D-Luciferin</td><td>Gold Biotechnology (St Louis, MO, USA)</td><td>LUCK-1G</td><td></td></tr><tr><td>Isoflurane</td><td>Henry Schein (Melville, NY, USA)</td><td>1182097</td><td></td></tr><tr><td>IVIS Lumina X5</td><td>PerkinElmer (Melville, NY, USA)</td><td>n/a</td><td>In vivo bioluminescence imaging system</td></tr><tr><td>J:NU mouse</td><td>The Jackson Laboratory (Bar Harbor, ME, USA)</td><td>007850</td><td>Athymic nude model</td></tr><tr><td>Kendall Hypoallergenic Paper Tape</td><td>Cardinal Health (Dublin, OH, USA)</td><td>1914C</td><td></td></tr><tr><td>Kimtech Surgical Nitrile Gloves</td><td>Kimberly-Clark Professional (Irving, TX, USA)</td><td>56892</td><td>Sterile Gloves</td></tr><tr><td>Nair Hair Remover Lotion</td><td>Church and Dwight (Trenton, NJ, USA)</td><td>n/a</td><td>Depilatory agent</td></tr><tr><td>NOD scid gamma (NSG) mouse</td><td>The Jackson Laboratory (Bar Harbor, ME, USA)</td><td>005557</td><td>Immunodeficient model</td></tr><tr><td>Phosphate-Buffered Saline (PBS), 1x</td><td>Corning (Corning, NY, USA)</td><td>21-040-CV</td><td></td></tr><tr><td>Puralube Vet Ointment</td><td>Med Vet International</td><td>PH-PURALUBE-VET</td><td>Eye ointment</td></tr><tr><td>Sheathes</td><td>Sheathing Technologies (Morgan Hill, CA, USA)</td><td>10040</td><td>Sterile Ultrasound Probe Covers</td></tr><tr><td>Sure-Seal Induction Chamber</td><td>Braintree Scientific (Braintree, MA, USA)</td><td>EZ-17 85</td><td>Anesthesia induction chamber</td></tr><tr><td>Transducer MX550D</td><td>FUJIFILM VisualSonics (Toronto, ON, Canada)</td><td>n/a</td><td>Vevo 3100 imaging probe (25-55 MHz, Centre Transmit: 40 MHz)</td></tr><tr><td>Trypan Blue, 0.4% solution in PBS</td><td>MP Biomedicals (Solon, OH, USA)</td><td>91691049</td><td></td></tr><tr><td>Vevo 3100 Imaging System</td><td>FUJIFILM VisualSonics (Toronto, ON, Canada)</td><td>n/a</td><td>Ultrasound imaging system</td></tr><tr><td>Vevo 3100 Lab Software</td><td>FUJIFILM VisualSonics (Toronto, ON, Canada)</td><td>n/a</td><td>Version 3.2.7 for imaging and analysis</td></tr><tr><td>Vevo Compact Dual Anesthesia System</td><td>FUJIFILM VisualSonics (Toronto, ON, Canada)</td><td>n/a</td><td>Tabletop isoflurane-based anesthesia unit</td></tr><tr><td>Vevo Imaging Station</td><td>FUJIFILM VisualSonics (Toronto, ON, Canada)</td><td>n/a</td><td>Procedural platform</td></tr></tbody></table>

a minimally invasive, accurate, and efficient technique for intrathymic injection in mice

Intrathymic injection in mouse models is an important technique for studying thymic and immune function, including genetic and acquired T cell disorders. This requires methods for the direct deposition of reagents and/or cells into the thymus of living mice. Traditional methods of intrathymic injection include thoracic surgery or minimally invasive percutaneous blind injections, both of which have significant limitations. Ultra-high frequency ultrasound imaging devices have made image-guided percutaneous injections possible in mice, greatly improving the injection accuracy of the percutaneous injection approach and enabling the injection of smaller targets. However, image-guided injections rely on the utilization of an integrated rail system, making this a rigid and time-consuming procedure. A unique, safe, and efficient method for percutaneous intrathymic injections in mice is presented here, eliminating reliance on the rail system for injections. The technique relies on using a high-resolution micro-ultrasound unit to image the mouse thymus noninvasively. Using a free-hand technique, a radiologist can place a needle tip directly into the mouse thymus under sonographic guidance. Mice are cleaned and anesthetized before imaging. For an experienced radiologist adept at ultrasound-guided procedures, the learning period for the stated technique is quite short, typically within one session. The method has a low morbidity and mortality rate for the mice and is much faster than current mechanically assisted techniques for percutaneous injection. It allows the investigator to efficiently perform precise and reliable percutaneous injections of thymuses of any size (including very small organs such as the thymus of aged or immunodeficient mice) with minimal stress on the animal. This method enables the injection of individual lobes if desired and facilitates large-scale experiments due to the time-saving nature of the procedure.

The successful implementation of this technique relies on a few key steps to be followed. First, reliable identification of the thymus gland itself has to be ensured. In young mice, this is straightforward due to the gland&#39;s large size (Figure 3A). In older mice or immunodeficient mice, it can be more challenging; however, it is still very feasible with modern ultrasound equipment (Figure 3B,C). Second, it is critically important to set the ...

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A Minimally Invasive, Accurate, and Efficient Technique for Intrathymic Injection in Mice

The present protocol describes an interventional radiology procedure established for intrathymic injection in mice to avoid the risk of open surgery and improve the accuracy of blind percutaneous injections.

Intrathymic injection in mouse models is an important technique for studying thymic and immune function, including genetic and acquired T cell disorders. ...

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2.3K Views.  Hackensack University Medical Center. The present protocol describes an interventional radiology procedure established for intrathymic injection in mice to avoid the risk of open surgery and improve the accuracy of blind percutaneous injections.

Video: A Minimally Invasive, Accurate, and Efficient Technique for Intrathymic Injection in Mice

Intralymphatic Immunotherapy and Vaccination in Mice

Vaccines are typically injected subcutaneously or intramuscularly for stimulation of immune responses. The success of this requires efficient drainage of vaccine to lymph nodes where antigen presenting cells can interact with lymphocytes for generation of the wanted immune responses. The strength and the type of immune responses induced also depend on the density or frequency of interactions as well as the microenvironment, especially the content of cytokines. As only a minute fraction of peripherally injected vaccines reaches the lymph nodes, vaccinations of mice and humans were performed by direct injection of vaccine into inguinal lymph nodes, i.e.&#160;intralymphatic injection. In man, the procedure is guided by ultrasound. In mice, a small (5-10 mm) incision is made in the inguinal region of anesthetized animals, the lymph node is localized and immobilized with forceps, and a volume of 10-20 &#956;l of the vaccine is injected under visual control. The incision is closed with a single stitch using surgical sutures. Mice were vaccinated with plasmid DNA, RNA, peptide, protein, particles, and bacteria as well as adjuvants, and strong improvement of immune responses against all type of vaccines was observed. The intralymphatic method of vaccination is especially appropriate in situations where conventional vaccination produces insufficient immunity or where the amount of available vaccine is limited.

Prophylactic and therapeutic vaccination often fails to stimulate strong immune responses due to week drainage of the vaccine to lymph nodes and consequently poor involvement of immune cells. By direct injection of vaccine to lymph nodes, so-called intralymphatic injection, vaccine efficacy can be strongly improved and vaccine doses can be reduced.

Vaccines are typically injected subcutaneously or intramuscularly for stimulation of immune responses. The success of this requires efficient drainage of ...

Ultrasound-guided Transthoracic Intramyocardial Injection in Mice

Murine models of cardiovascular disease are important for investigating pathophysiological mechanisms and exploring potential regenerative therapies. Experiments involving myocardial injection are currently performed by direct surgical access through a thoracotomy. While convenient when performed at the time of another experimental manipulation such as coronary artery ligation, the need for an invasive procedure for intramyocardial delivery limits potential experimental designs. With ever improving ultrasound resolution and advanced noninvasive imaging modalities, it is now feasible to routinely perform ultrasound-guided, percutaneous intramyocardial injection. This modality efficiently and reliably delivers agents to a targeted region of myocardium. Advantages of this technique include the avoidance of surgical morbidity, the facility to target regions of myocardium selectively under ultrasound guidance, and the opportunity to deliver injectate to the myocardium at multiple, predetermined time intervals. With practiced technique, complications from intramyocardial injection are rare, and mice quickly return to normal activity on recovery from anesthetic. Following the steps outlined in this protocol, the operator with basic echocardiography experience can quickly become competent in this versatile, minimally invasive technique.

Echocardiography-guided percutaneous intramyocardial injection represents an efficient, reliable, and targetable modality for the delivery of gene transfer agents or cells into the murine heart. Following the steps outlined in this protocol, the operator can quickly become competent in this versatile, minimally invasive technique.

Murine models of cardiovascular disease are important for investigating pathophysiological mechanisms and exploring potential regenerative therapies. ...

Medicine

Minimally Invasive Cisterna Magna Injection Model for Leptomeningeal Metastasis Studies in Mice

Leptomeningeal metastasis (LM), the spread of cancer cells into the cerebrospinal fluid (CSF)-filled leptomeninges, is a rare yet devastating complication of advanced solid tumors. Patients with LM often have a poor prognosis, with survival measured in weeks to months. Developing in vivo models that accurately replicate the complexities of LM is essential for understanding its cellular and pathological mechanisms and evaluating potential therapies. Murine LM models are typically created through intra-cardiac, carotid artery, or cisterna magna injection of tumor cells. However, intra-cardiac or carotid injections often result in substantial extracranial and brain tumor burden, complicating bioluminescent imaging and leading to mortality unrelated to LM. Meanwhile, conventional cisterna magna injection requires invasive procedures, such as skin incision and muscle dissection, making it both traumatic and resource intensive. Here, we describe a minimally invasive procedure for tumor cell injection into the leptomeningeal space through the cisterna magna without the need for a skin incision. This approach reduces extracranial tumor formation, minimizes surgical trauma, and shortens the time and postoperative care required compared to other surgical methods. Importantly, it consistently induces LM with minimal brain parenchyma infiltration, as confirmed by two-photon microscopy and histological analysis. This streamlined approach offers an efficient and reliable model for studying LM in preclinical research.

The protocol describes a method for injecting tumor cells through the percutaneous puncture route into cisterna magna that robustly induces leptomeningeal metastasis in mice, reducing trauma and extracranial tumor burden.

Leptomeningeal metastasis (LM), the spread of cancer cells into the cerebrospinal fluid (CSF)-filled leptomeninges, is a rare yet devastating complication ...

Cancer Research

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter implantation. The technical limitation of this surgical technique lies in the finesse of the hands. The injection is rapid, especially for a trained experimenter, and since tissue disruption with this technique is minimal, repeated injections are possible; moreover immune reaction to foreign tools (e.g. catheter) does not occur, thereby giving a better and more specific read out of spinal cord modulation. Since the application of the substance is largely limited to the target region of the spinal cord, drugs do not need to be applied in large dosages, and more importantly unwanted effects on other tissue, as observed with a systemic delivery, could be circumvented1,2. Moreover, we combine this technique with in vivo transfection of nucleic acid with the help of polyethylenimine (PEI) reagent3, which provides tremendous versatility for studying spinal functions via delivery of pharmacological agents as well as gene, RNA, and protein modulators.

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a simple and rapid technique of intrathecal needle puncture for a localized transfection of siRNA in the lumbar spinal cord in mouse under short lasting light anesthesia.

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter ...

Biology

Intravenous Injections in Neonatal Mice

Intravenous injection is a clinically applicable manner to deliver therapeutics. For adult rodents and larger animals, intravenous injections are technically feasible and routine. However, some mouse models can have early onset of disease with a rapid progression that makes administration of potential therapies difficult. The temporal (or facial) vein is just anterior to the ear bud in mice and is clearly visible for the first two days after birth on either side of the head using a dissecting microscope. During this window, the temporal vein can be injected with volumes up to 50 &#956;l. The injection is safe and well tolerated by both the pups and the dams. A typical injection procedure is completed within 1-2 min, after which the pup is returned to the home cage. By the third postnatal day the vein is difficult to visualize and the injection procedure becomes technically unreliable. This technique has been used for delivery of adeno-associated virus (AAV) vectors, which in turn can provide almost body-wide, stable transgene expression for the life of the animal depending on the viral serotype chosen.

Animal models of pediatric disease can experience early onset and aggressive disease progression. Clinically relevant therapy delivery to young mouse models can be technically difficult. This protocol describes a non-invasive intravenous injection method for newborn mice within the first two postnatal days of life.

Intravenous injection is a clinically applicable manner to deliver therapeutics. For adult rodents and larger animals, intravenous injections are ...

Performing Intracerebroventricular Injection in Mice Using a Free-Hand Approach

Free-Hand Intracerebroventricular Injections in Mice

The investigation of neuroendocrine systems often requires the delivery of drugs, viruses, or other experimental agents directly into the brains of mice. An intracerebroventricular (ICV) injection allows the widespread delivery of the experimental agent throughout the brain (particularly in the structures near the ventricles). Here, methods for making free-hand ICV injections in adult mice are described. By using visual and tactile landmarks on the heads of mice, injections into the lateral ventricles can be made rapidly and reliably. The injections are made with a glass syringe held in the experimenter&#39;s hand and placed at approximate distances from the landmarks. Thus, this technique does not require a stereotaxic frame. Furthermore, this technique requires only brief isoflurane anesthesia, which permits the subsequent assessment of mouse behavior and/or physiology in awake, freely behaving mice. Free-hand ICV injection is a powerful tool for the efficient delivery of experimental agents into the brains of living mice and can be combined with other techniques such as frequent blood sampling, neural circuit manipulation, or in vivo recording to investigate neuroendocrine processes.

Author Spotlight: Insights, Innovations, and Future Avenues in Reproductive Neurocontrol 

Here, a simple and rapid approach for performing intracerebroventricular injections in mice using a free-hand approach (that is, without a stereotaxic device) is described.

The investigation of neuroendocrine systems often requires the delivery of drugs, viruses, or other experimental agents directly into the brains of mice. ...

Neuroscience

A Comprehensive Protocol for Delivering Intrathecal Injections in Neonatal Mice

Intrathecal Injection of Newborn Mouse for Genome Editing and Drug Delivery

Intrathecal injection is a commonly employed procedure in both pediatric and adult clinics, serving as an effective means to administer medications and treatments. By directly delivering medications and treatments into the cerebrospinal fluid of the central nervous system, this method achieves higher localized drug concentrations while reducing systemic side-effects compared to other routes such as intravenous, subcutaneous, or intramuscular injections. Its importance extends beyond clinical settings, as intrathecal injection plays a vital role in preclinical studies focused on treating neurogenetic disorders in rodents and other large animals, including non-human primates. However, despite its widespread application, intrathecal injection in young, particularly neonatal pups, poses significant technical challenges due to their small size and fragile nature. Successful and reliable administration of intrathecal injections in newborn mice requires meticulous attention to detail and careful consideration of various factors. Thus, there is a crucial need for a standardized protocol that not only provides instructions but also highlights key technical considerations and good laboratory practices to ensure procedural consistency, as well as the safety and welfare of the animals.
To address this unmet need, we present a detailed and comprehensive protocol for performing intrathecal injections specifically in newborn pups on postnatal day 1 (P1). By following the step-by-step instructions, researchers can confidently perform intrathecal injections in neonatal pups, enabling the accurate delivery of drugs, antisense oligos, and viruses for gene replacement or genome editing-based treatments. Furthermore, the importance of adhering to good laboratory practices is emphasized to maintain the well-being of animals and ensure reliable experimental outcomes. This protocol aims to address the technical challenges associated with intrathecal injections in neonatal mice, ultimately facilitating advances in the field of neurogenetic research that aims to develop potential therapeutic interventions.

Author Spotlight: Intrathecal Injection &#8211; An Efficient and Reliable Delivery Method to Test the Efficacy of Gene Editing in Neonatal Mouse Brains

The present protocol outlines step-by-step instructions for performing intrathecal injections in neonatal mice for gene editing and drug delivery.

Intrathecal injection is a commonly employed procedure in both pediatric and adult clinics, serving as an effective means to administer medications and ...

A Simple and Efficient Method for In Vivo Cardiac-specific Gene Manipulation by Intramyocardial Injection in Mice

Gene manipulation specifically in the heart significantly potentiate the investigation of cardiac disease pathomechanisms and their therapeutic potential. In vivo cardiac-specific gene delivery is commonly achieved by either systemic or local delivery. Systemic injection via tail vein is easy and efficient in manipulating cardiac gene expression by using recombinant adeno-associated virus 9 (AAV9). However, this method requires a relatively high amount of vector for efficient transduction, and may result in nontarget organ gene transduction. Here, we describe a simple, efficient, and time-saving method of intramyocardial injection for in vivo cardiac-specific gene manipulation in mice. Under anesthesia (without ventilation), the pectoral major and minor muscles were bluntly dissected, and the mouse heart was quickly exposed by manual externalization through a small incision at the fourth intercostal space. Subsequently, adenovirus encoding luciferase (Luc) and vitamin D receptor (VDR), or short hairpin RNA (shRNA) targeting VDR, was injected with a Hamilton syringe into the myocardium. Subsequent in vivo imaging demonstrated that luciferase was successfully overexpressed specifically in the heart. Moreover, Western blot analysis confirmed the successful overexpression or silencing of VDR in the mouse heart. Once mastered, this technique can be used for gene manipulation, as well as injection of cells or other materials such as nanogels in the mouse heart.

A Simple and Efficient Method for In Vivo Cardiac-specific Gene Manipulation by Intramyocardial Injection in Mice

Here we present a protocol for cardiac-specific gene manipulation in mice. Under anesthesia, the mouse hearts were externalized through the fourth intercostal space. Subsequently, adenoviruses encoding specific genes were injected with a syringe into the myocardium, followed by protein expression measurement via in vivo imaging and Western blot analysis.

Gene manipulation specifically in the heart significantly potentiate the investigation of cardiac disease pathomechanisms and their therapeutic potential. ...

Intrafemoral Injection for Bone Marrow Study in Live Mice

Simplified Intrafemoral Injections Using Live Mice Allow for Continuous Bone Marrow Analysis

Despite the complexity of hematopoietic cell transplantation in humans, researchers commonly perform intravenous or intrafemoral (IF) injections in mice. In murine models, this technique has been adapted to enhance the seeding efficiency of transplanted hematopoietic stem and progenitor cells (HSPCs). This paper describes a detailed step-by-step technical procedure of IF injection and the following bone marrow (BM) aspiration in mice that allows for serial characterization of cells present in the BM. This method enables the transplantation of valuable samples with low cell numbers that are particularly difficult to engraft by intravenous injection. This procedure facilitates the creation of xenografts that are critical for pathological analysis. While it is easier to access peripheral blood (PB), the cellular composition of PB does not reflect the BM, which is the niche for HSPCs. Therefore, procedures providing access to the BM compartment are essential for studying hematopoiesis. IF injection and serial BM aspiration, as described here, allow for the prospective retrieval and characterization of cells enriched in the BM, such as HSPCs, without sacrificing the mice.

This protocol describes the intrafemoral injection of a few hematopoietic or leukemic stem cells, including gene-edited cells, in murine xenograft models, which will not only enable the quick and safe transplantation of cells but also serial analyses of the bone marrow.

Despite the complexity of hematopoietic cell transplantation in humans, researchers commonly perform intravenous or intrafemoral (IF) injections in mice. ...

Compound Administration IV

Source: Kay Stewart, RVT, RLATG, CMAR; Valerie A. Schroeder, RVT, RLATG. University of Notre Dame, IN
There are many commonly used routes for compound administration in laboratory mice and rats. Protocols may, however, require the use of the less commonly used routes: intracardiac, footpad, and retro-orbital injections. Specialized training is essential for these procedures to be performed successfully. Justification for these routes may need to be provided to gain Institutional Animal Care and Use Committee (IACUC) approval.

Compound Administration in Rodents- Specialized Injections

Source: Kay Stewart, RVT, RLATG, CMAR; Valerie A. Schroeder, RVT, RLATG. University of Notre Dame, IN
There are many commonly used routes for compound ...

A Minimally Invasive, Accurate, and Efficient Technique for Intrathymic Injection in Mice

Summary

Explore More Videos

Intralymphatic Immunotherapy and Vaccination in Mice

Ultrasound-guided Transthoracic Intramyocardial Injection in Mice

Minimally Invasive Cisterna Magna Injection Model for Leptomeningeal Metastasis Studies in Mice

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

Intravenous Injections in Neonatal Mice

Free-Hand Intracerebroventricular Injections in Mice

Intrathecal Injection of Newborn Mouse for Genome Editing and Drug Delivery

A Simple and Efficient Method for In Vivo Cardiac-specific Gene Manipulation by Intramyocardial Injection in Mice

Simplified Intrafemoral Injections Using Live Mice Allow for Continuous Bone Marrow Analysis

Compound Administration IV