This work was supported by Dorothy D. Radford Endowment fund from the West Virginia University Mary Babb Randolph Cancer Center.&#160; The use of the West Virginia University Animal Models and Imaging Facility (AMIF) and Microscopy Imaging Facility (MIF) (supported by the Mary Babb Randolph Cancer Center and NIH grants P20 RR16440, P30 RR032138/GM103488 and S10 RR026378) is gratefully acknowledged.

All animal procedures demonstrated in this protocol have been reviewed and approved by the West Virginia University Animal Care and Use Committee under protocols 11-0412 and 14-0514 and conducted in accordance with the principles and procedures outlined in the NIH Guide for the Care and Use of Animals.
1. Animal Preparation
<ol>
	<li>Anesthetize a single mouse in an induction chamber using 3% isofluorane mixed with 1.5 L/min 100% oxygen. Remove the animal from the induction chamber and place in a supine position on the imaging platform preheated to 40 &#176;C and maintained between 37-42 &#176;C (Figure 2A). Confirm anesthesia by the lack of response to a toe pinch.</li>
	<li>Position the mouse snout within the nosecone connected to the anesthesia system. Apply anesthesia to maintain steady state sedation (1.5% isofluorane mixed with 1.5 L/min 100% oxygen).</li>
	<li>Apply eye lubricant the each eye to prevent drying. Apply electrode gel to the electrodes and use tape to secure each of the four paws to the corresponding electrode. The electrode pads will transmit the animal&#8217;s ECG to the imaging system to allow monitoring of heart rate and respiration rate. Lubricate and insert the rectal temperature probe for continuous monitoring of body temperature. Normal mouse body temperature is 36.9 &#176;C. 1-2 &#176;C variation is normal while under anesthesia.</li>
	<li>Use depilatory cream to remove the fur from the neck of the mouse. Rinse the neck region with water-soaked gauze to remove hair and excess depilatory cream. Optionally, use additional application of depilatory cream to remove any remaining body hair (Figure 2B).</li>
</ol>
2. Identification and Image Acquisition of Mouse Cervical Lymph Nodes Using HFUS
<ol>
	<li>To begin, apply a layer of warmed ultrasound gel to the neck area devoid of fur. Use liberal application of gel for optimal image quality (Figure 2C). Avoid introducing air bubbles in the gel during application, which can interfere with ultrasonic imaging.</li>
	<li>Adjust the imaging platform 20-30&#176; so the mouse is positioned with the head slightly elevated. This position helps to ensure the optimal respiration rate for the mouse. Place the 40 MHz transducer transversely in the mounting system and carefully lower it until the front of the transducer scanhead is immersed in the ultrasound gel (Figure 2C). 
	NOTE: Be sure to not put excessive pressure onto the mouse neck, as it could cause undue respiratory distress. In addition, it is helpful for imaging to have a buffer of gel between the transducer and the mouse.</li>
</ol>
B-Mode Imaging and Detection of Lymph Nodes:
<ol start="3">
	<li>Using the computer controlling the HFUS acquisition software, adjust the brightness (B-) mode settings to the following parameters: Gain 22 dB, depth 10.00 mm, width 14.08 mm. 
	NOTE: These settings are a suggested starting point, and may require slight adjusting for optimal image acquisition between different applications. Observe normal cervical lymph nodes as oval hypoechoic structures near the skin surface within a surrounding hyperechoic field. The appearance of diseased lymph nodes can vary between models. To systematically image all lymph nodes within the neck region use the following steps:
	<ol>
		<li>Use the Y-axis to scan the neck in a cranial to caudal manner towards the thoracic region. Use X-axis to center the image.</li>
		<li>Identify major landmarks: buccal cavity, tongue, and thyroid gland (Figures 3A, B and C; respectively); tilt the imaging platform to horizontally adjust the ventral surface of the mouse neck, making both sides of the neck appear even in the B-mode image. The amount of tilting depends upon the physiology of each individual mouse.</li>
	</ol>
	</li>
	<li>Conduct a 3D scan of the entire neck region from the buccal cavity/tongue region to the thyroid gland, in order to map lymph nodes and associated landmarks throughout the neck.
	<ol>
		<li>Locate the tongue/buccal cavity region (Figure 3A) and note the numerical location on the Y scale.</li>
		<li>Locate thyroid gland (Figure 3C) and note the numerical location on the Y scale.</li>
		<li>Calculate the difference between the obtained values in (2.4.1) and (2.4.2) to determine the total length in mm for the imaged neck region.</li>
		<li>Use the Y knob to center the transducer on the midpoint of the determined total length.</li>
		<li>Press &#8220;3D&#8221;. Enter the total length. For 3D step size, use 0.076 mm to acquire the image series stack for the entire neck region.</li>
	</ol>
	</li>
	<li>Once scanning is complete, select the right or left side of the neck and center the transducer on an individual lymph node of interest, then raise the 40 MHz transducer off the mouse. Remove the 40 MHz transducer and replace with a 50 MHz microscan transducer (also in a transverse position) to obtain higher-resolution images. Replenish the ultrasound gel on the mouse neck and lower the 50 MHz transducer into the ultrasound gel.</li>
</ol>
3D Power Doppler Imaging
<ol start="6">
	<li>Conduct 3D scans with power Doppler to asses volume and vascularity of individual cervical lymph nodes as follows:
	<ol>
		<li>Press the Power button on the system keyboard to acquire power Dopper and adjust the following power-mode settings: PRF 4 KHz, Doppler gain 34 dB, 2D gain 30 dB, depth 5.00 mm, width 4.73 mm. NOTE: As before, these settings are a suggested starting point and may be modified as needed for optimal image acquisition in various models.</li>
		<li>Locate the cranial-most point of the lymph node of interest and note the location on the Y scale.</li>
		<li>Locate the most caudal point of the same node and note the location on the Y scale.</li>
		<li>Calculate the distance difference to determine the total length of the lymph node (see step 2.4.3).</li>
		<li>Center the transducer on the midpoint of the determined total length, using the Y-scale location.</li>
		<li>Press &#8220;3D&#8221; and enter total lymph node length. Use 0.051 mm for the step size. 
		NOTE: Due to the proximity of the transducer to the chest region, the 50 MHz transducer may result in an unstable Doppler image due to detection of normal respiratory motion. This can be eliminated by using the &#8220;Respiration Gating&#8221; option available under the &#8220;Physiological&#8221; tab.</li>
		<li>Surround the selected lymph node with the yellow box that designates the region to be analyzed by power Doppler and select &#8220;3D scan&#8221; to acquire images. Raise the transducer off of the mouse and move it to the opposite side of the neck. Lower the transducer onto the mouse and repeat the steps described above to image lymph nodes on this side of the neck.</li>
	</ol>
	</li>
	<li>Save image sets for subsequent analysis.</li>
</ol>
3. Cervical Lymph Node Biopsy
<ol>
	<li>Select the desired lymph node for biopsy and maintain HFUS imaging with the 50 MHz transducer. Chose the largest visible cervical lymph node in each side of the mouse neck. Lymph node enlargement typically indicates an inflammatory response, and therefore such nodes are ideal candidates for biopsy. 
	NOTE: We have found it is very difficult to conduct biopsies on cervical nodes smaller than 10 mm3.</li>
	<li>Prepare the needle and syringe for biopsy by placing a 1 ml syringe with an attached 27 G, 0.5 inch needle into the syringe holder. Adjust the needle holder to orient the needle 90&#176; to the mouse neck (Figure 4A).</li>
	<li>Prepare by elevating the entire mouse platform to the level of the needle. Achieve this by removing the 3D motor and switching to a taller platform, or by placing a solid object of suitable height under the provided short platform. Use a plastic microfuge tube rack for this purpose. If necessary, rotate the platform 180&#176; in order to biopsy nodes located on the side of the neck opposite to the needle apparatus.</li>
	<li>Adjust the acquisition settings by selecting &#8220;Preferences&#8221;, then by choosing &#8220;Max &#38; Extended buffer&#8221;. Enlarge the field of view to a depth of 8.00 mm and width of 9.73 mm. Turn on the needle guide using the Screen Keys dial. The needle guide will predict the path of the needle on the screen and allows the user to line up the lymph node of interest in the correct location for biopsy.</li>
	<li>Ensure that the lymph node remains constantly in view by centering the lymph node in the middle or slightly to the left of center in the screen (Figure 4B). To obtain the entire cine loop of the procedure, press Pre-trigger on the system keyboard before beginning the biopsy.</li>
	<li>Adjust the needle holder until the needle tip comes into view and contacts the skin (Figure 4B). Advance the needle with a firm, fast push to puncture the skin. Continue to advance the needle until the tip also punctures the capsule (Figure 4C) and is visible within the medulla (Figure 4D).</li>
	<li>Once the needle is properly located within the node, gently pull the syringe plunger back between the 200-300 &#181;l demarcations to conduct the biopsy (Figure 4D and 4E). Note that biopsy material is typically not visible within the syringe.</li>
	<li>Gently remove the needle from the mouse neck. Expel the syringe contents into a 1.5 ml microfuge tube. Remove the needle from the syringe, leaving the needle in the tube. Collect 1 ml of biopsy media (from an aliquot, separate from the stock source) with the same syringe, and then reattach the needle to the syringe while keeping the needle in the tube.
	<ol>
		<li>Rinse the syringe and needle with the biopsy media by expelling the biopsy media into the tube. 
		NOTE: Do not pull back on the plunger while the needle is attached at any point after the biopsy. This reduces the risk of losing the biopsy material due to the small sample size.</li>
	</ol>
	</li>
	<li>Confirm lymph node content by histological means (Figure 4F) and analyze by additional methods (histochemistry, flow cytometry, etc.) as appropriate.</li>
	<li>Once biopsy is complete, turn off the anesthesia and remove the rectal temperature probe. Remove excess ultrasound gel from mouse with gauze and remove the tape from each paw.</li>
	<li>Remove the mouse from the imaging platform and return to a cage. Minimal bleeding from biopsy site may occur, but this stops without intervention. Monitor the mouse during recovery until full activity is resumed.</li>
</ol>
4. Image Analysis of Cervical Lymph Nodes
<ol>
	<li>In the ultrasound software, select the image for analysis and navigate to the &#8220;Image Processing&#8221; tab. Choose &#8220;Load into 3D&#8221;.</li>
	<li>Select &#8220;3D Reconstructed Image&#8221; in top left corner, clicking on the &#8220;Display Single Pane&#8221; button. Use the zoom function to enlarge the image if desired. Toggle &#8220;Display Layout&#8221; to view the image only in B-mode, which removes the power Doppler overlay from view. This makes it easier to see the edges of the lymph node during subsequent 3-D analysis. Scroll through the image series to locate the beginning of the lymph node.</li>
	<li>To circumscribe the lymph node, navigate to the &#8220;3D Settings&#8221; tab. Select &#8220;volume&#8221;, then the &#8220;Start&#8221; button next to &#8220;Parallel&#8221;.</li>
	<li>Draw contours around the area of interest within individual images by scrolling. Continue until images that encompass the entire lymph node are marked. Choose &#8220;Finish&#8221; to complete the analysis.</li>
	<li>At the bottom of the image, 3D volume and % vascularity will be automatically displayed. 
	NOTE: 3D volume corresponds to the lymph node volume, and percent vascularity represents the percentage of the lymph node positive for blood flow by power Doppler.</li>
	<li>Toggle &#8220;Display Layout&#8221; to view the power Doppler imaging as an overlay on the B- mode image. On the surface view observe a net view of the volume area of interest. Export the images in tagged image file (TIF) format or 3D scans as movies (.avi) for further use.</li>
</ol>

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Publication costs for this article are sponsored by Visual Sonics.

The described protocol allows for the visualization and in situ evaluation of murine cervical lymph nodes using non-invasive HFUS imaging. The use of B-mode and power Doppler imaging to visualize cervical lymph node morphology, volume and lymph node blood flow provides for an experimental analysis of preclinical mouse model systems similar to that used for the characterization of cervical patient nodes in clinical practice. The ability to monitor the cervial lymph nodes via fine needle biopsy also provides a useful technique for detecting immune cell alterations and the presence of foreign cell types or bacteria during oral cavity disease-induced lymphadenopathies in mice. The ease of use and low cost associated with HFUS allows for the rapid screening of cervical lymph node status in a wide variety of animal models.
A critical step in this protocol is the initial successful identification of the cervical lymph nodes in HFUS images. Our facility has an assortment of HFUS transducers as described, so we have used them to obtain the highest quality images. However, if the transducers we describe are unavailable, it is possible to adapt the imaging using other transducers.&#160; For this purpose, adjusting image depth and width to obtain an adequate image is all that is required. Resolution of such images may vary, but it should still be possible to obtain high quality images by using HFUS. Imaging landmarks of the oral cavity and the thyroid gland will greatly aid in orienting the user to the proper region where the lymph nodes are localized. The characteristic oval-shaped, hypoechoic nature and superficial location close to the skin surface allows for rapid confirmatory identification of the cervical lymph nodes within the proper neck region. While all three nodes may be visible in a single imaging plane (Figure 3B), typically one or two nodes are captured during imaging. Minor adjustments of the transducer position can be conducted to render different imaging planes visible, allowing visualization of all nodes on a single side of the neck.
While we have found the described technique reliable for identifying cervical lymph nodes, there are specific limitations to the imaging and biopsy technique. The superficial nature of mouse cervical lymph nodes confers excessive mobility when even slight pressure is applied to the skin via the transducer head.&#160; This can be counteracted by slowly applying the transducer head into the ultrasound gel on the mouse neck until the landmark images are identified. Lymph node mobility can also complicate the fine needle biopsy, especially when using transducers in the higher resolution (50 MHz) range. Centered images of lymph nodes for biopsy are typically pushed out of the field of view due to the force of the biopsy needle needed to puncture the overlying skin and capsule. This can be remedied by the off-center positioning of the lymph node towards the direction of needle entry, providing space for the lymph node to be pushed across but still remain within the field of view during biopsy. In our experience, lymph nodes &#62;10 mm3 are very difficult to biopsy, and are often pushed by the needle rather than penetrated during needle advancement. Thus, biopsy is best reserved for enlarged lymph nodes where the size is &#62;10 mm3 to ensure sufficient node target size and stability within the cervical region. In addition, biopsy material may not contain sufficient cell numbers for procedures where larger cell numbers are required (e.g., flow cytometry). &#160;
HFUS has been used to successfully visualize orthotopic HNSCC tumors25, and has the potential to monitor cervical node metastasis in mice with oral tumors24. In addition to ultrasound, bioluminescence imaging has also been used to visualize orthotopic oral tumor formation and cervical lymph node metastasis in live mice26,27. As an alternative approach, bioluminescence imaging has a distinct advantage over HFUS in being able to directly quantify tumor progression and metastatic burden over time in the same animal. While undeniably useful, bioluminescence imaging is unable to measure many of the parameters visualized by HFUS, including lymph node morphology, nodal volumes or blood flow. Bioluminescence imaging also requires specialized dark boxes to maintain mice during imaging, rendering this technique unsuitable for adaptation for fine needle biopsy.
Furthermore, bioluminescence imaging requires production of tumor cells that stably express the luciferase enzyme, allowing this technique to be used only in cases of orthotopic xenografts with luciferase-transfected tumor cells in immunocompromised mice, or with inducible tissue-specific transgenic systems that restrict luciferase expression&#160; in&#160; a&#160; spatio-temporal&#160; manner&#160; specific to the tissue of tumor origin. In contrast, HFUS can be used in conjunction with bioluminescence images in these models, as well as being capable of imaging cervical&#160; lymph nodes in models of carcinogen-induced oral tumors in mice with complete immune systems28,29. While HFUS may be more adaptable to most mouse models of oral cancer, the combined information that can be obtained from bioluminescence and HFUS imaging in systems where tumor cells express luciferase can provide a more complete picture of cervical lymph node involvement than either imaging modality alone.
The ability to identify and detect mouse cervical lymph nodes in real time allows for this technique to be used in most models of oral disease that result in inflammatory lymphadenopathy where the animal can be maintained in an inverted position under short-term anesthesia. Detection of lymph node metastasis or bacterial infection and the concomitant impact on lymph node morphology in living animals presents a significant benefit over traditional methods that require lymph nodes to be removed from dead animals for histological processing. Combining HFUS with fine needle biopsy allows a means for conducting routine pathological analysis of cervical lymph nodes, similar to what is conducted in the clinic, providing an improved method for monitoring disease progression in most current mouse models of oral cavity diseases.

Lymph drainage of interstitial tissue fluid is the main method of dissemination for infectious microorganisms and cancers arising in the oral and maxillofacial region1,2. Clinical evaluation of cervical lymph nodes is a common diagnostic practice used to determine the presence or progression of diseases that originate in the oral cavity. This underscores the importance of the collecting cervical lymph nodes as valuable anatomic sites for oral disease diagnosis3. Several specialized imaging methodologies are clinically utilized for identifying diseased cervical lymph nodes. These include positron emission tomography (PET), computed tomography (CT) and magnetic resonance imaging (MR). While highly valuable, these methods all require extensive patient preparation, highly specialized equipment and/or chemical infusion into the circulation to enable or enhance the imaging process.
Sonographic imaging (ultrasound; US) is a commonly applied technique used to image cervical lymph nodes presenting with lymphadenopathy due to infection or metastatic involvement4-6. US is often combined with PET-CT and MR imaging to provide a comprehensive representation of patient lymph node status, helping to determine tumor staging and necessity for surgical excision7. The non-invasive nature of US also has inherent advantages over other imaging modalities, including ease of use, low cost, minimal patient discomfort and preparation. The superficial subcutaneous location of most cervical lymph nodes allows for US to guide minimally invasive fine needle aspiration biopsies with increased precision, improving diagnostic accuracy8.
Commercial high-frequency (HF) US provides detailed resolution of internal anatomic structures to 30 &#181;m9. Using transducers ranging from 22-70 MHz, HFUS has been readily applied to a variety of experimental rodent systems to allow real-time imaging of internal organs in vivo.&#160; HFUS has been adapted for visualization of tumor formation in conventional brightness (B)-mode, as well as with a number of general and specialized contrast enhancement agents9. Using power Doppler with HFUS provides the ability to monitor blood flow within mouse tumors, allowing a comprehensive assessment of angiogenic perfusion in live mice10,11. HFUS has been used to visualize diseased mouse lymph nodes within the main body cavity, demonstrating parallel utility of this technology to clinical practice. In particular, inflammatory and metastatic visceral lymph node alterations have been observed in mouse models of cancer harboring breast12,13, pancreatic14, colorectal15 and lung16 tumors, as well as fibrous histocytomas17, and an aged mouse model of acquired hydronephrosis18. These examples solidify the value of HFUS as a powerful investigative tool for tumor-induced lymphadenopathy in a wide variety of rodent systems. &#160;
Several models of bacterial infection19,20 and head and neck squamous cell carcinoma (HNSCC)21,22 have been developed to study these diseases in the preclinical setting. In contrast to humans, mice contain three cervical lymph nodes that survey lymph from oral cavity tissues (mandibular, submandibular mandibular and superficial parotid23). Recently, we reported the use of HFUS to map the location and morphology of these lymph nodes, monitoring&#160; changes in lymph&#160; node volume and blood flow in a carcinogen-induced mouse model of HNSCC24. Here, we provide a detailed protocol for the use of HFUS for identifying, imaging and analyzing cervical lymph nodes in living mice. This protocol also demonstrates the feasibility of using HFUS to conduct image- guided fine needle biopsy of enlarged mouse cervical lymph nodes, allowing histological monitoring of changes in cervical lymph node content and pathologies over time in the same animal. This protocol is readily adaptable to allow for the detailed study of cervical lymph node pathologies resultant from any invasive oral cavity disease in mice.

<table><tbody><tr><td>Vevo2100 High Resolution Micro-ultrasound Imaging System, with integrated software Version 1.6.0</td><td>VisualSonics, Toronto, Ontario, Canada</td><td>VS-11945</td><td></td></tr><tr><td>Power Dopper Mode and 3D Mode</td><td>VisualSonics, Toronto, Ontario, Canada</td><td>VS-11952; VS-11484</td><td></td></tr><tr><td>Vevo compact anesthesia system</td><td>VisualSonics, Toronto, Ontario, Canada</td><td></td><td></td></tr><tr><td>Vevo integrated rail system including 3D motor and micromanipulator for injections</td><td>VisualSonics, Toronto, Ontario, Canada</td><td>SA-11983</td><td></td></tr><tr><td>Thermasonic Gel Warmer</td><td>VisualSonics, Toronto, Ontario, Canada</td><td></td><td>Optional</td></tr><tr><td>Transducers &amp;ndash; MS-550D (Broadband frequency: 22 MHz - 55 MHz)</td><td>VisualSonics, Toronto, Ontario, Canada</td><td>VS-11960</td><td>Referred to as 40 MHz Transducer</td></tr><tr><td>Transducers &amp;ndash; MS-700 (Broadband frequency: 30 MHz - 70 MHz)</td><td>VisualSonics, Toronto, Ontario, Canada</td><td>VS-12026</td><td>Referred to as 50 MHz Transducer</td></tr><tr><td>Ophthalmic Ointment</td><td>Patterson Veterinary</td><td>07-888-1663</td><td></td></tr><tr><td>Electrode gel</td><td>Parker Laboratories</td><td>174-1525</td><td></td></tr><tr><td>Tape</td><td>Medical Arts Press</td><td>174-153000</td><td></td></tr><tr><td>Depilatory Cream</td><td>Carter Products</td><td></td><td></td></tr><tr><td>Cotton swabs</td><td>General Supply</td><td></td><td></td></tr><tr><td>Gauze</td><td>Fisher Scientific</td><td>22-037-907</td><td></td></tr><tr><td>Water</td><td>General Supply</td><td></td><td></td></tr><tr><td>Lubricating gel</td><td>Parker Laboratories</td><td>57-05</td><td></td></tr><tr><td>Ultrasound gel</td><td>Parker Laboratories</td><td>01-50</td><td></td></tr><tr><td>Microcentrifuge tube rack</td><td>General Supply</td><td></td><td>Used to raise mouse platform for optimal biopsy position</td></tr><tr><td>27 G &amp;frac12; inch needle with 1 ml syringe</td><td>Fisher Scientific</td><td>14-826-87</td><td></td></tr><tr><td>ThinPrep PreservCyt Solution</td><td>Hologic</td><td>70097-002</td><td>Refered to as biopsy media</td></tr><tr><td>Microcentrifuge tubes</td><td>General Supply</td><td></td><td></td></tr><tr><td>Thinprep 2000 processor</td><td>Cytyc, Marlborough, MA</td><td></td><td>Blue Filter</td></tr><tr><td>Olympus AX70 Provis Microscope</td><td>Olympus, Center Valley, PA</td><td></td><td></td></tr></tbody></table>

high-frequency ultrasound imaging of mouse cervical lymph nodes

High-frequency ultrasound (HFUS) is widely employed as a non-invasive method for imaging internal anatomic structures in experimental small animal systems. HFUS has the ability to detect structures as small as 30 &#181;m, a property that has been utilized for visualizing superficial lymph nodes in rodents in brightness (B)-mode. Combining power Doppler with B-mode imaging allows for measuring circulatory blood flow within lymph nodes and other organs. While HFUS has been utilized for lymph node imaging in a number of mouse&#160; model systems, a detailed protocol describing HFUS imaging and characterization of the cervical lymph nodes in mice has not been reported. Here, we show that HFUS can be adapted to detect and characterize cervical lymph nodes in mice. Combined B-mode and power Doppler imaging can be used to detect increases in blood flow in immunologically-enlarged cervical nodes. We also describe the use of B-mode imaging to conduct fine needle biopsies of cervical lymph nodes to retrieve lymph tissue for histological&#160; analysis. Finally, software-aided steps are described to calculate changes in lymph node volume and to visualize changes in lymph node morphology following image reconstruction. The ability to visually monitor changes in cervical lymph node biology over time provides a simple and powerful technique for the non-invasive monitoring of cervical lymph node alterations in preclinical mouse models of oral cavity disease.

The overall schematic for the imaging and biopsy procedures is shown in Figure 1. The key steps in the procedure include proper preparation of the mouse for imaging, identification of the cervical lymph nodes, correct preparation and conducting of the needle biopsy, and analysis of B-mode and Doppler images to measure volume and the amount of vascularity within each selected node using computer software.
HFUS imaging of mouse cervical lymph nodes requires applying and maintaining proper anesthesia throughout the imaging period (Figure 2A), as well as complete removal of the hair covering the entire neck area (Figure 2B). The liberal application of ultrasound gel to the depilated region ensures a clear HFUS signal during the procedure (Figure 2C).
HFUS imaging of the neck region is aided by the visualization of cervical anatomical landmarks that produce characteristic sonographic images. Figure 3 shows examples of the key organs (Figure 3A-C), cervical lymph nodes in B-mode (Figure 3D) and in power Doppler mode (Figure 3E).
Real-time HFUS imaging in anesthetized mice allows for guided fine needle biopsy of cervical nodes similar to what is conducted in clinical practice. Placement of the biopsy needle and attached collection syringe to the controlling microinjector equipment is shown in Figure 4A. Subsequent B-mode sonographic images show ideal needle placement prior to biopsy (Figure 4B), needle tip entry into a cervical lymph node (Figure 4C), and needle position during biopsy (Figure 4D). Close-up image shows the needle tip within the medulla of the lymph node (Figure 4E). Processing of the biopsy components by cytospin reveals abundant lymphoid cell clusters and associated connective tissue, verifying successful lymph node biopsy (Figure 4F).
Computational-based analysis of HFUS images allows for detailed information to be obtained regarding lymph node architecture, volume and vascular flow. Using power Doppler mode and 3D volume measurements, percent vascularity (PV) can be calculated from image series encompassing entire nodes (Figure 5A). Additionally, 3D imaging allows for virtual lymph node reconstruction, revealing overall lymph node topography (Figure 5B).
<img alt="Mouse imaging, biopsy, and analysis workflow; ultrasound, biopsy, vascularity assessment steps." src="/files/ftp_upload/52718/52718fig1.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Overview schematic of the steps involved in diagnostic HFUS cervical lymph node imaging in mice. The key steps include 1: Preparing the mice for HFUS imaging and obtaining 40 and 50 MHz resolution images of the neck region containing the three mouse cervical lymph nodes. 2: Fine needle image-guided biopsy of cervical lymph nodes and subsequent histological analysis of biopsied material. 3: Computer-aided image analysis and 3D reconstruction of lymph node images obtained in B-mode and Doppler to determine the respective lymph node volume and percent (%) of vascular flow.
<img alt="Mouse imaging experiment setup with 3D MS and MI, for anatomical study, equipment in use." src="/files/ftp_upload/52718/52718fig2.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Overview of the high resolution in vivo micro-imaging system for cervical lymph node assessment and biopsy. (A) The HFUS system is shown with an anesthetized mouse prepared for cervical lymph node imaging. Also shown is the microinjector (MI) and 3D-motor stage (3D MS) accessory equipment. (B) Close up view of an anesthetized mouse prepared for HFUS imaging with hair removed in neck region. (C) The same mouse with the 50 MHz transducer in place on the neck. Note the extra ultrasound gel used to facilitate neck region imaging.
<img alt="Ultrasound imaging of buccal cavity anatomy; diagrams highlight tongue, mandibular, thyroid, glands." src="/files/ftp_upload/52718/52718fig3.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3: Representative HFUS cervical anatomy images in B-mode and power Doppler. (A, B) B-mode images of the oral cavity, showing the buccal cavity (BC) and tongue (T) visualized by imaging closest to the nasal cavity. The three cervical lymph nodes found on each side of the neck (labeled M, mandibular; SM, submandibular; SP, superficial parotid), appear as a group of hypoechoic structures in a single imaging plane as shown (B). (C)The thyroid gland (Th) as visualized in the upper thoracic region, appearing as a solid, echogenic butterfly-shaped structure. (A-C) were visualized with a 40 MHz transducer; scale bar = 1 mm. (D, E) Representative images of normal (D) and enlarged (E) cervical lymph nodes with B-mode and power Doppler (red). Dotted lines outline individual lymph nodes. Scale bar = 0.5 mm.
<img alt="Ultrasound-guided biopsy process in diagram: needle positioning, entry, biopsy, cytospin result." src="/files/ftp_upload/52718/52718fig4.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4: Cervical lymph node biopsy set-up, imaging and cytospin analysis of biopsy material. (A) The imaging platform showing the micro-injector and needle placement near the mouse neck. A wide microcentrifuge tube rack (orange block) is used to slightly raise the platform, enabling proper needle placement while still allowing space for the 3D-motor stage. This arrangement minimizes time spent removing the motor stage for each mouse. (B-D) Whole neck HFUS images taken from a video of a cervical lymph node biopsy using the 50 MHz transducer. (B) HFUS B-mode image showing the needle positioned to the side of the neck prior to biopsy. The needle tip is the hyperechoic structure just below the position of the needle guide (green dotted line) superimposed during imaging to denote the needle trajectory. The lymph node is in the center of the image. Scale bar = 1 mm. (C) Needle entry into the lymph node. (D) Biopsy of the cervical lymph node. (E) Zoomed biopsy of cervical lymph node. Scale bar = 0.5 mm. (F) Cytospin analysis of representative biopsy lymph material confirming successful biopsy. Scale bar = 100 µm.
<img alt="Ultrasound imaging diagram; panel A shows tissue boundary, panel B shows 3D volume reconstruction." src="/files/ftp_upload/52718/52718fig5.jpg" data-altupdatedat="1747911167000" data-alt="Figure 5"> 
Figure 5: Computer analysis of 3D cervical lymph node images. (A) Representative screenshot of a lymph node analyzed using computer software. The node is circumscribed in blue; analysis results show 3D Volume and percent vascularity (PV) as indicated. (B) A surface view image of the same node after 3D analysis. Renders entire volume of lymph node based on measurements taken in A.

Watch this Scientific Journal Video about High-frequency Ultrasound Imaging of Mouse Cervical Lymph Nodes at JoVE.com

High-frequency Ultrasound Imaging of Mouse Cervical Lymph Nodes

This protocol describes the application of high-frequency ultrasound (HFUS) for imaging mouse cervical lymph nodes. This technique optimizes visualization and quantification of cervical lymph node morphology, volume and blood flow. Image-guided biopsy of cervical lymph nodes and processing of lymph tissue for histological evaluation is also demonstrated.

High-frequency ultrasound (HFUS) is widely employed as a non-invasive method for imaging internal anatomic structures in experimental small animal ...

high-frequency-ultrasound-imaging-of-mouse-cervical-lymph-nodes

Research

JoVE Journal

Medicine

19.0K Views.  West Virginia University.  Dit protocol beschrijft de toepassing van hoogfrequente ultrageluid (HFUS) voor imaging muizen cervicale lymfeknopen. Deze techniek optimaliseert visualisatie en kwantificatie van cervicale lymfeknopen morfologie, volume en de bloedstroom. Beeld-geleide biopsie van cervicale lymfeklieren en verwerking van lymfeweefsel voor histologische evaluatie wordt ook aangetoond. 

Video: High-frequency Ultrasound Imaging of Mouse Cervical Lymph Nodes

Dissection and 2-Photon Imaging of Peripheral Lymph Nodes in Mice

Two-photon imaging has revealed an elegant choreography of motility and cellular interactions within the lymph node under basal conditions and at the initiation of an immune response 1. Here, we present methods for adoptive transfer of labeled T cells, isolation of lymph nodes, and imaging motility of CD4+ T cells in the explanted lymph node as first described in 2002 2. Two-photon imaging of immune cells requires that the cells are fluorescently labeled, either by staining with a cell tracker dye or by expressing a fluorescent protein. We demonstrate the adoptive transfer procedure of injecting cells derived from donor mice into the tail vein of a recipient animal, where they home to lymphoid organs within approximately 15-30 min. We illustrate the isolation of a lymph node and describe methods to ensure proper mounting of the excised lymph node. Other considerations such as proper oxygenation of perfused media, temperature, and laser power are discussed. Finally, we present 3D video images of naive CD4+ T cells exhibiting steady state motility at 37&deg;C.

Two-photon imaging has uncovered lymphocyte motility and cellular interactions within the lymph node under basal conditions and durring an immune response 1. Here, we demonstrate adoptive transfer of T cells, isolation of lymph nodes, and imaging motility of CD4+ T cells in the explanted lymph node.

Two-photon imaging has revealed an elegant choreography of motility and cellular interactions within the lymph node under basal conditions and at the ...

Biology

Murine Superficial Lymph Node Surgery

In the field of immunology, to understand the progression of an immune response against a vaccine, an infection or a tumour, the response is often followed over time. Similarly, the study of lymphocyte homeostasis requires time course experiments. Performing these studies within the same mouse is ideal to reduce the experimental variability as well as the number of mice used. Blood withdrawal allows performance of time course experiments, but it only gives information about circulating lymphocytes and provides a limited number of cells1-4. Since lymphocytes circulating through the body and residing in the lymph nodes have different properties, it is important to examine both locations. The sequential removal of lymph nodes by surgery provides a unique opportunity to follow an immune response or immune cell expansion in the same mouse over time. Furthermore, this technique yields between 1-2x106 cells per lymph node which is sufficient to perform phenotypic characterization and/or functional assays. Sequential lymph node surgery or lymphadenectomy has been successfully used by us and others5-11. Here, we describe how the brachial and inguinal lymph nodes can be removed by making a small incision in the skin of an anesthetised mouse. Since the surgery is superficial and done rapidly, the mouse recovers very quickly, heals well and does not experience excessive pain. Every second day, it is possible to harvest one or two lymph nodes allowing for time course experiments. This technique is thus suitable to study the characteristics of lymph node-residing lymphocytes over time. This approach is suitable to various experimental designs and we believe that many laboratories would benefit from performing sequential lymph node surgeries.

To follow the progression of an immune response over time within the same mouse, lymph nodes can be sequentially removed by surgery. Here, we describe how this technique can be performed.

In the field of immunology, to understand the progression of an immune response against a vaccine, an infection or a tumour, the response is often ...

Immunology and Infection

Intravital Imaging of the Mouse Popliteal Lymph Node

Lymph nodes (LNs) are secondary lymphoid organs, which are strategically located throughout the body to allow for trapping and presentation of foreign antigens from peripheral tissues to prime the adaptive immune response. Juxtaposed between innate and adaptive immune responses, the LN is an ideal site to study immune cell interactions1,2. Lymphocytes (T cells, B cells and NK cells), dendritic cells (DCs), and macrophages comprise the bulk of bone marrow-derived cellular elements of the LN. These cells are strategically positioned in the LN to allow efficient surveillance of self antigens and potential foreign antigens3-5. The process by which lymphocytes successfully encounter cognate antigens is a subject of intense investigation in recent years, and involves an integration of molecular contacts including antigen receptors, adhesion molecules, chemokines, and stromal structures such as the fibro-reticular network2,6-12. 
Prior to the development of high-resolution real-time fluorescent in vivo imaging, investigators relied on static imaging, which only offers answers regarding morphology, position, and architecture. While these questions are fundamental in our understanding of immune cell behavior, the limitations intrinsic with this technique does not permit analysis to decipher lymphocyte trafficking and environmental clues that affect dynamic cell behavior. Recently, the development of intravital two-photon laser scanning microscopy (2P-LSM) has allowed investigators to view the dynamic movements and interactions of individual cells within live LNs in situ12-16. In particular, we and others have applied this technique to image cellular behavior and interactions within the popliteal LN, where its compact, dense nature offers the advantage of multiplex data acquisition over a large tissue area with diverse tissue sub-structures11,17-18. It is important to note that this technique offers added benefits over explanted tissue imaging techniques, which require disruption of blood, lymph flow, and ultimately the cellular dynamics of the system. Additionally, explanted tissues have a very limited window of time in which the tissue remains viable for imaging after explant. With proper hydration and monitoring of the animal's environmental conditions, the imaging time can be significantly extended with this intravital technique. Here, we present a detailed method of preparing mouse popliteal LN for the purpose of performing intravital imaging.

Recent advances in 2-photon microscopy have enabled real-time in situ imaging of live tissues in animal models, thereby enhancing our ability to investigate cellular behavior in both physiologic and pathologic conditions. Here, we outline the preparations required to perform intravital imaging of the mouse popliteal lymph node.

Lymph nodes (LNs) are secondary lymphoid organs, which are strategically located throughout the body to allow for trapping and presentation of foreign ...

Non-invasive Optical Imaging of the Lymphatic Vasculature of a Mouse

The lymphatic vascular system is an important component of the circulatory system that maintains fluid homeostasis, provides immune surveillance, and mediates fat absorption in the gut. Yet despite its critical function, there is comparatively little understanding of how the lymphatic system adapts to serve these functions in health and disease1. Recently, we have demonstrated the ability to dynamically image lymphatic architecture and lymph "pumping" action in normal human subjects as well as in persons suffering lymphatic dysfunction using trace administration of a near-infrared fluorescent (NIRF) dye and a custom, Gen III-intensified imaging system2-4. NIRF imaging showed dramatic changes in lymphatic architecture and function with human disease. It remains unclear how these changes occur and new animal models are being developed to elucidate their genetic and molecular basis. In this protocol, we present NIRF lymphatic, small animal imaging5,6 using indocyanine green (ICG), a dye that has been used for 50 years in humans7, and a NIRF dye-labeled cyclic albumin binding domain (cABD-IRDye800) peptide that preferentially binds mouse and human albumin8. Approximately 5.5 times brighter than ICG, cABD-IRDye800 has a similar lymphatic clearance profile and can be injected in smaller doses than ICG to achieve sufficient NIRF signals for imaging8. Because both cABD-IRDye800 and ICG bind to albumin in the interstitial space8, they both may depict active protein transport into and within the lymphatics. Intradermal (ID) injections (5-50 &mu;l) of ICG (645 &mu;M) or cABD-IRDye800 (200 &mu;M) in saline are administered to the dorsal aspect of each hind paw and/or the left and right side of the base of the tail of an isoflurane-anesthetized mouse. The resulting dye concentration in the animal is 83-1,250 &mu;g/kg for ICG or 113-1,700 &mu;g/kg for cABD-IRDye800. Immediately following injections, functional lymphatic imaging is conducted for up to 1 hr using a customized, small animal NIRF imaging system. Whole animal spatial resolution can depict fluorescent lymphatic vessels of 100 microns or less, and images of structures up to 3 cm in depth can be acquired9. Images are acquired using V++ software and analyzed using ImageJ or MATLAB software. During analysis, consecutive regions of interest (ROIs) encompassing the entire vessel diameter are drawn along a given lymph vessel. The dimensions for each ROI are kept constant for a given vessel and NIRF intensity is measured for each ROI to quantitatively assess "packets" of lymph moving through vessels.

Recently developed imaging techniques using near-infrared fluorescence (NIRF) may help elucidate the role the lymphatic system plays in cancer metastasis, immune response, wound repair, and other lymphatic-associated diseases.

The lymphatic  vascular system is an important component of the circulatory system that  maintains fluid homeostasis, provides immune surveillance, and ...

Highly Resolved Intravital Striped-illumination Microscopy of Germinal Centers

Monitoring cellular communication by intravital deep-tissue multi-photon microscopy is the key for understanding the fate of immune cells within thick tissue samples and organs in health and disease. By controlling the scanning pattern in multi-photon microscopy and applying appropriate numerical algorithms, we developed a striped-illumination approach, which enabled us to achieve 3-fold better axial resolution and improved signal-to-noise ratio, i.e. contrast, in more than 100 &#181;m tissue depth within highly scattering tissue of lymphoid organs as compared to standard multi-photon microscopy. The acquisition speed as well as photobleaching and photodamage effects were similar to standard photo-multiplier-based technique, whereas the imaging depth was slightly lower due to the use of field detectors. By using the striped-illumination approach, we are able to observe the dynamics of immune complex deposits on secondary follicular dendritic cells &#8211; on the level of a few protein molecules in germinal centers.

High-resolution intravital imaging with enhanced contrast up to 120 &#181;m depth in lymph nodes of adult mice is achieved by spatially modulating the excitation pattern of a multi-focal two-photon microscope. In 100 &#181;m depth we measured resolutions of 487 nm (lateral) and 551 nm (axial), thus circumventing scattering and diffraction limits.

Monitoring cellular communication by intravital deep-tissue multi-photon microscopy is the key for understanding the fate of immune cells within thick ...

Analysis of Lymph Node Volume by Ultra-High-Frequency Ultrasound Imaging in the Braf/Pten Genetically Engineered Mouse Model of Melanoma

Tyr::CreER+,BrafCA/+,Ptenlox/lox&#160;genetically engineered mice (Braf/Pten mice) are widely used as an in vivo model of metastatic melanoma. Once a primary tumor has been induced by tamoxifen treatment, an increase in metastatic burden is observed within 4-6 weeks after induction. This paper shows how Ultra-High-Frequency UltraSound (UHFUS) imaging can be exploited to monitor the increase in metastatic involvement of the inguinal lymph nodes by measuring the increase in their volume.
The UHFUS system is used to scan anesthetized mice with a UHFUS linear probe (22-55 MHz, axial resolution 40 &#181;m). B-mode images from the inguinal lymph nodes (both left and right sides) are acquired in a short-axis view, positioning the animals in dorsal recumbency. Ultrasound records are acquired using a 44 &#181;m step size on a motorized mechanical arm. Afterward, two-dimensional (2D) B-mode acquisitions are imported into the software platform for ultrasound image post-processing, and inguinal lymph nodes are identified and segmented semi-automatically in the acquired cross-sectional 2D images. Finally, a total reconstruction of the three-dimensional (3D) volume is automatically obtained along with the rendering of the lymph node volume, which is also expressed as an absolute measurement.
This non-invasive in vivo technique is very well tolerated and allows the scheduling of multiple imaging sessions on the same experimental animal over 2 weeks. It is, therefore, ideal to assess the impact of pharmacological treatment on metastatic disease.

Melanoma is a very aggressive disease that quickly spreads to other organs. This protocol describes the application of ultra-high-frequency ultrasound imaging, coupled with 3D rendering, to monitor the volume of the inguinal lymph nodes in the Braf/Pten mouse model of metastatic melanoma.

Tyr::CreER+,BrafCA/+,Ptenlox/lox&#160;genetically engineered mice (Braf/Pten mice) are widely used as an in vivo model of metastatic melanoma. Once a ...

Cancer Research

High-Resolution Ultrasonography for the Analysis of Orthotopic ATC Tumors in a Genetically Engineered Mouse Model

Anaplastic thyroid carcinoma (ATC) is associated with a poor prognosis and short median survival time, but no effective treatment improves the outcomes significantly. Genetically engineered murine models that mimic ATC&#39;s progression may help researchers to study treatments for this disease. Crossing three different genotypes of mice, a TPO-cre/ERT2; BrafCA/wt; Trp53&#916;ex2-10/&#916;ex2-10&#160;transgenic ATC model was developed. The ATC murine model was induced by an intraperitoneal injection of tamoxifen with overexpression of BrafV600E and deletion of Trp53, and the tumors were generated within about 1 month. High-resolution ultrasound was applied to investigate the tumor initiation and progression, and the dynamic growth curve was obtained by measuring the tumor sizes. Compared to magnetic resonance imaging (MRI) and computed tomography scanning, ultrasound has advantages in observing the ATC murine model, such as being noninvasive, portable, in real-time, and without radiation exposure. High-resolution ultrasound is suitable for dynamic and multiple measurements. However, ultrasonographic examination of the thyroid in mice requires relevant anatomical knowledge and experience. This article provides a detailed procedure for utilizing high-resolution ultrasound to scan tumors in the transgenic ATC model. Meanwhile, ultrasonic parameter adjustment, ultrasound scanning skills, anesthesia and recovery of the animals, and other elements that need attention during the process are listed.

The present protocol describes high-frequency ultrasonography for visualizing the entire mouse thyroid gland and monitoring the growth of anaplastic thyroid carcinoma.

Anaplastic thyroid carcinoma (ATC) is associated with a poor prognosis and short median survival time, but no effective treatment improves the outcomes ...

Non-Invasive Ultrasound Assessment of Endometrial Cancer Progression in Pax8-Directed Deletion of the Tumor Suppressors Arid1a and Pten in Mice

Uterine cancers can be studied in mice due to the ease of handling and genetic manipulation in these models. However, these studies are often limited to assessing pathology post-mortem in animals euthanized at multiple time points in different cohorts, which increases the number of mice needed for a study. Imaging mice in longitudinal studies can track the progression of disease in individual animals, reducing the number of mice needed. Advances in ultrasound technology have allowed for the detection of micrometer-level changes in tissues. Ultrasound has been used to study follicle maturation in ovaries and xenograft growth but has not been applied to morphological changes in the mouse uterus. This protocol examines the juxtaposition of pathology with in vivo imaging comparisons in an induced endometrial cancer mouse model. The features observed by ultrasound were consistent with the degree of change seen by gross pathology and histology. Ultrasound was found to be highly predictive of the observed pathology, supporting the incorporation of ultrasonography into longitudinal studies of uterine diseases such as cancer in mice.

Non-Invasive Ultrasound Assessment of Endometrial Cancer Progression in Pax8-Directed Deletion of the Tumor Suppressors Arid1a and Pten in Mice

This protocol describes a method for monitoring the progression of morphological changes over time in the uterus in an inducible mouse model of endometrial cancer using ultrasound imaging with correlation to gross and histological changes.

Uterine cancers can be studied in mice due to the ease of handling and genetic manipulation in these models. However, these studies are often limited to ...

Visualizing Lymph Node Structure and Cellular Localization using Ex-Vivo Confocal Microscopy

Lymph nodes (LNs) are organs spread within the body, where the innate immune responses can connect with the adaptive immunity. In fact, LNs are strategically interposed in the path of the lymphatic vessels, allowing intimate contact of tissue antigens with all resident immune cells in the LN. Thus, understanding the cellular composition, distribution, location and interaction using ex vivo whole LN imaging will add to the knowledge on how the body coordinates local and systemic immune responses. This protocol shows an ex vivo imaging strategy following an in vivo administration of fluorescent-labeled antibodies that allows a very reproducible and easy-to-perform methodology by using conventional confocal microscopes and stock reagents. Through subcutaneous injection of antibodies, it is possible to label different cell populations in draining LNs without affecting tissue structures that can be potentially damaged by a conventional immunofluorescence microscopy technique.

This protocol describes a technique to image different cell populations in draining lymph nodes without alterations in the organ structure.

Lymph nodes (LNs) are organs spread within the body, where the innate immune responses can connect with the adaptive immunity. In fact, LNs are ...

Ultrafast Doppler Ultrasound for Hepatic Microvasculature Analysis in Mice

Imaging and Quantification of the Hepatic Vasculature of Mice Using Ultrafast Doppler Ultrasound

Non-invasive in vivo imaging of the vasculature is a powerful tool for studying disease mechanisms in rodents. To achieve high sensitivity imaging of the microvasculature using Doppler ultrasound methods, imaging modalities employing the concept of ultrafast imaging are preferred. By increasing the frame rate of the ultrasound scanner to thousands of frames per second, it becomes possible to improve the sensitivity of the blood flow down to 2 mm/s and to obtain functional information about the microcirculation in comparison to a sensitivity of around 1 cm/s in conventional Doppler modes. While Ultrafast Doppler ultrasound (UFUS) imaging has become adopted in neuroscience, where it can capture brain activity through neurovascular coupling, it presents greater challenges when imaging the vasculature of abdominal organs due to larger motions linked to breathing. The liver, positioned anatomically under the diaphragm, is particularly susceptible to out-of-plane movement and oscillating respiratory motion. These artifacts not only adversely affect Doppler imaging but also complicate the anatomical analysis of vascular structures and the computation of vascular parameters. Here, we present a qualitative and quantitative imaging analysis of the hepatic vasculature in mice by UFUS. We identify major anatomical vascular structures and provide graphical illustrations of the hepatic macroscopical anatomy, comparing it to an in-depth anatomical assessment of the hepatic vasculature based on Doppler readouts. Additionally, we have developed a quantification protocol for robust measurements of hepatic blood volume of the microvasculature over time. To contemplate further research, qualitative vascular analysis provides a comprehensive overview and suggests a standardized terminology for researchers working with mouse models of liver disease. Furthermore, it offers the opportunity to apply ultrasound as a non-invasive complementary method to inspect hepatic vascular defects in vivo and measure functional microvascular alterations deep within the organ before unraveling blood vessel anomalies at the micron scale levels using ex vivo staining on tissue sections.

Ultrafast Doppler ultrasound (UFUS) with high spatial resolution (100 &#956;m) and sensitivity represents a suitable noninvasive imaging modality for obtaining a comprehensive qualitative overview of the hepatic vasculature. UFUS also facilitates quantitative measurements of the microvasculature, aiming to enhance our understanding of vascular disease mechanisms.

Non-invasive in vivo imaging of the vasculature is a powerful tool for studying disease mechanisms in rodents. To achieve high sensitivity imaging of the ...

High-frequency Ultrasound Imaging of Mouse Cervical Lymph Nodes

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