This work was supported by the NIH (R43-DK126607, TJC, TLK, MFR) and the Mayo Foundation.

All steps in this protocol comply with the Mayo Clinic (Rochester, MN) animals use guidelines and have been approved by the Mayo Clinic Institutional Animal Care and Use Committee.
1. Animal model
<ol>
	<li>Obtain mice from commercial sources. 
	&#8203;NOTE: C57BL/6J&#160;(age = 18 months; n = 22 (7 males, 15 females)) (Table of Materials) and Pkd1(RC/RC) (age = 18 months; n = 9 (3 males, 6 females))4 were used for the study.</li>
</ol>
2. Hair removal
<ol>
	<li>Anesthetize the animal using vaporized isoflurane (2%-3%) in an anesthesia chamber. Ensure that the animal is sufficiently anesthetized by performing a toe pinch. Ensure that the mouse is breathing at a rate of approximately 60-90 breaths/min without obvious air gulping. 
	NOTE: Vaporized isoflurane was chosen due to the ability to control and fine-tune the animal&#39;s level of anesthesia.&#160;Isoflurane mixed with 100% oxygen was utilized.</li>
	<li>Use an electric razor to shave the desired skin area for US measurements. 
	NOTE: In this study, the focus is on kidney and cardiac measurements, so the animal&#39;s chest and back were shaved.</li>
	<li>Apply the depilatory cream to the animal&#39;s shaved skin with a cotton-tipped applicator. Wipe the cream off after 30-60 s using a dry paper towel. Then, continue to wipe away depilatory cream residue using wet gauze pads. Ensure the hair removal is complete prior to beginning scans. 
	&#8203;NOTE: Any remaining hair will interfere with the image quality; complete hair removal is crucial. However, it is necessary to wipe the depilatory cream off within a minute to prevent injuring the animal&#39;s skin. Leftover depilatory cream on the animal can stain the imaging membrane of the US machine. Ensure proper cleaning of the animal using wet gauze pads.</li>
</ol>
3. Animal positioning
<ol>
	<li>Turn on the US machine and the switch for the internal heating element for the US bays. Turn on the appropriate anesthesia valves for the corresponding US bays that will be used. 
	NOTE: The US machine must be turned on prior to starting the imaging US software.</li>
	<li>Fill the bays with enough water to cover the membrane. Place the anesthetized animal in the nose cone.
	<ol>
		<li>For cardiac measurements, place the animal in a prone position and angle it approximately 30&#176; to the right.</li>
		<li>For kidney measurements, place the animal in a straight and supine position. Gently flatten the animal&#39;s body to minimize the air pockets between the animal and the membrane (Figure 1B). 
		NOTE: Running a finger alongside the animal&#39;s body can remove any air bubbles adhering to the skin, which will interfere with imaging. Use water or US saline gel as US medium; but water facilitates the rapid imaging discussed here.</li>
	</ol>
	</li>
</ol>
4. Ultrasound measurements
<ol>
	<li>In the imaging US software, select US Acquisition and allow the instrument to initialize.</li>
	<li>Kidney Imaging using B-mode.
	<ol>
		<li>Enter a name for the study in the appropriate dialog box.</li>
		<li>Enter the animal ID in the appropriate dialog box. 
		&#8203;NOTE: Up to three animals can be imaged sequentially.</li>
		<li>Click on the tab labeled 3D to initialize 3D scanning.</li>
		<li>Using the live camera feed, move the region of interest (ROI) box to contain the kidneys. 
		NOTE: The size of the ROI may be adjusted depending on the size of the image target. Increasing the size of the ROI will slightly increase the scan time (Figure 1C). Depending on ROI size, the scanner will automatically adjust the number of passes required for the transducer to scan the entire ROI.</li>
		<li>Depending on the type of image desired, select Linear Array or Wobbler, to choose the transducer. Select from Fast Scan or Optimized for either transducer. 
		NOTE: Linear array is faster and usually provides adequate imaging. The wobbler transducer can be utilized if unable to acquire the desired image quality with the linear array. The wobbler transducer utilizes a higher frequency wavelength and therefore provides more definition in images. The Fast Scan option acquires medium quality images with no breathing correction at a higher speed. While the Optimized option requires slightly more time, it provides better image quality and allows for breathing correction. This manuscript will focus on using the Optimized linear array preset.</li>
		<li>Click on the Live View to see a live image of the desired viewing area.</li>
		<li>To move the transducer, click on the circular Go To button, and then click on the area of the mouse to be imaged. Use the directional arrows with predetermined step sizes to finely adjust the transducer position.</li>
		<li>Click on the arrow at the side of the Control Panel window to open the Settings Tray. Click on Scan Protocol to open scanning settings.</li>
		<li>Determine the ideal focal depth and adjust the sliders for the TGC (time gain compensation) to change the image contrast to best view the kidneys. After achieving the desired settings, click on the arrow again to close the Settings Tray.</li>
		<li>Press the Acquire Static button to commence 3D scanning. 
		NOTE: Progress of the scan is indicated via a progress bar in the bottom left of the software.</li>
		<li>Once the scanning is complete, check the image quality by clicking on the blue download arrow to open the scan in Preview Mode. Move through the 3D frames of the scan by dragging the red arrow at the top of the window. To zoom in or out, hold the Ctrl key and use the scroll button on the mouse.</li>
		<li>To scan additional animals, refresh the animal ID dialog boxes, input the new IDs, and then repeat the above steps. 
		NOTE: When the scans are complete, dry the animals off and return them to their home cages.</li>
	</ol>
	</li>
	<li>Cardiac imaging using M mode. 
	NOTE: M-mode is the display of an ultrasound wave along a specific, chosen ultrasound line.
	<ol>
		<li>Click on the M Mode tab to activate cardiac imaging. 
		&#8203;NOTE: Only one animal can be imaged at a time.</li>
		<li>Ensure that the animal is in the appropriate position for cardiac scans (step 2.2). Move the ROI to the approximate location of the heart.</li>
		<li>Push the Heart Finder button to locate the heart. Use the heat map to optimize the location of the transducer. Use the circular Go To button to move the transducer to the left ventricle (Figure 2A).</li>
		<li>Go to the Live View and use the arrow buttons to place the transducer between the papillary muscles (Figure 2B). To verify the proper placement of the transducer, view the live M Mode trace (Figure 2D).</li>
		<li>Once the transducer is in the appropriate position, click on Acquire Sequence. Click on the blue download arrow to view the completed scan.</li>
	</ol>
	</li>
</ol>
5. Kidney analysis (mechanics of analysis)
<ol>
	<li>Open the Image Analysis tab and select the Load Volume(s) button to input the image files/folders.
	<ol>
		<li>Segmentation
		<ol>
			<li>Move the colored lines representing various planes to find the kidney in all the planes.</li>
			<li>Click on Add to add a new segmentation. Provide a markup name for the file.</li>
			<li>Use the Surface Cut button to add points around the outer edge of the kidney in multiple planes. Click on Apply to generate segmentation through the entire kidney (Figure 3A).</li>
			<li>Scroll through the different frames in each plane and edit any inaccurate regions by clicking on Edit and dragging the points to the correct location.</li>
			<li>Alternatively, if the kidney is irregular in shape, use the Draw function to create an irregularly shaped trace around the kidney. Either click points around the edge or hold the left click to draw the trace freehand.</li>
			<li>To complete the trace with the Draw button, right-click with the mouse. Scroll ahead multiple frames and repeat the tracing process.</li>
			<li>Repeat until the frames are traced throughout the entire region of interest.</li>
			<li>If the Draw function is used, select Fill Between Slices to check and adjust the trace. Click on Initialize to see a preview. If the preview is acceptable, click on Apply. If not, click on Cancel and modify the tracings. 
			NOTE: The Erase button will allow deletion of a region within a trace. The Smooth button smooths the jagged edges of a trace.</li>
			<li>When image analysis is satisfactory, select Save to save the analysis to the original image file.</li>
		</ol>
		</li>
		<li>Measurement
		<ol>
			<li>Use the Calipers function to measure features within the image. Click on the Calipers button and left click the two points of the distance to be measured (Figure 3B).</li>
			<li>Save the measurement by clicking the disk button at the top of the screen.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
6. Cardiac analysis
<ol>
	<li>Select Load Volumes to input the file(s) to analyze.</li>
	<li>Select the Add option.</li>
	<li>Use the Point Placement tool to trace the systole and diastole for each layer of heart tissue. Continue with the other layers of heart tissue. Use the Edit function to modify the traces (Figure 2C,D). 
	NOTE: If the M-mode trace is very clear, use the Add (auto) button to automatically generate tracings of each layer.</li>
	<li>Select Apply to calculate the cardiac parameters. View the Quantify tab and select Export Statistics to export all data as a *.csv file (Figure 2E).</li>
</ol>

<ol><li>Cunha, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Preclinical+imaging%3A+an+essential+ally+in+modern+biosciences%5BTitle%5D)+AND+%22Molecular+Diagnostics+%26+Therapy%22%5BJournal%5D)">Preclinical imaging: an essential ally in modern biosciences.</a> Molecular Diagnostics & Therapy. 18 (2), 153-173 (2014).</li>
<li>Ultrasound. NIBIB. , Available from: 
 <a target="_blank" href="https://www.nibib.nih.gov/science-education/science-topics/ultrasound#pid-936">https://www.nibib.nih.gov/science-education/science-topics/ultrasound#pid-936</a> (2016).</li>
<li>Beaumont, N. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Assessing+polycystic+kidney+disease+in+rodents%3A+Comparison+of+robotic+3d+ultrasound+and+magnetic+resonance+imaging%5BTitle%5D)+AND+%22Kidney360%22%5BJournal%5D)">Assessing polycystic kidney disease in rodents: Comparison of robotic 3d ultrasound and magnetic resonance imaging.</a> Kidney360. 1 (10), 1128-1136 (2020).</li>
<li>Hopp, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Functional+polycystin-1+dosage+governs+autosomal+dominant+polycystic+kidney+disease+severity%5BTitle%5D)+AND+%22The+Journal+of+Clinical+Investigation%22%5BJournal%5D)">Functional polycystin-1 dosage governs autosomal dominant polycystic kidney disease severity.</a> The Journal of Clinical Investigation. 122 (11), 4257-4273 (2012).</li>
<li>Weiss, R. M., Ohashi, M., Miller, J. D., Young, S. G., Heistad, D. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Calcific+aortic+valve+stenosis+in+old+hypercholesterolemic+mice%5BTitle%5D)+AND+%22Circulation%22%5BJournal%5D)">Calcific aortic valve stenosis in old hypercholesterolemic mice.</a> Circulation. 114 (19), 2065-2069 (2006).</li>
<li>Czernuszewicz, T. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+new+preclinical+ultrasound+platform+for+widefield+3D+imaging+of+rodents%5BTitle%5D)+AND+%22Review+of+Scientific+Instruments%22%5BJournal%5D)">A new preclinical ultrasound platform for widefield 3D imaging of rodents.</a> Review of Scientific Instruments. 89 (7), 075107(2018).</li>
<li>Chong, W. K., Papadopoulou, V., Dayton, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Imaging+with+ultrasound+contrast+agents%3A+current+status+and+future%5BTitle%5D)+AND+%22Abdominal+Radiology+%28NY%29%22%5BJournal%5D)">Imaging with ultrasound contrast agents: current status and future.</a> Abdominal Radiology (NY). 43 (4), 762-772 (2018).</li>
<li>Nyankima, A. G., Kasoji, S., Cianciolo, R., Dayton, P. A., Chang, E. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Histological+and+blood+chemistry+examination+of+the+rodent+kidney+after+exposure+to+flash-replenishment+ultrasound+contrast+imaging%5BTitle%5D)+AND+%22Ultrasonics%22%5BJournal%5D)">Histological and blood chemistry examination of the rodent kidney after exposure to flash-replenishment ultrasound contrast imaging.</a> Ultrasonics. 98, 1-6 (2019).</li>
<li>Rojas, J. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ultrasound+measurement+of+vascular+density+to+evaluate+response+to+anti-angiogenic+therapy+in+renal+cell+carcinoma%5BTitle%5D)+AND+%22IEEE+Transactions+on+Biomedical+Engineering%22%5BJournal%5D)">Ultrasound measurement of vascular density to evaluate response to anti-angiogenic therapy in renal cell carcinoma.</a> IEEE Transactions on Biomedical Engineering. 66 (3), 873-880 (2019).</li>
<li>Boron, W. F., Boulpaep, E. L. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aBoron%2C+WF+%22Medical+Physiology.+3rd+edn%22">Medical Physiology. 3rd edn</a>. , Elsevier. (2017).</li>
</ol>

Several authors are either employed by, have a significant financial interest in, or are co-inventors on patents licensed by SonoVol, Inc. (CJM, RCG, TJC).

Ultrasound utilizes sound waves, and any barriers to sound wave propagation will interfere with image quality. Thus, complete hair removal of the area to be imaged is critical. It is also important to ensure complete removal of depilatory cream as it can cause burns/irritation of the animal&#39;s skin and discolor the transparent membrane of the scanner. Adequate water levels in the bays are necessary for optimal sound wave propagation, thus required for obtaining the highest image resolution. However, when the animal is in a prone position, ensure the animal&#39;s snout is above the water level or the animal is at risk of water inhalation. Optimization of imaging parameters, especially focal depth, is critical for obtaining high-quality images. Modifications to parameters may be necessary for individual animals.
Robotic US provides many advantages over traditional hand-held US modalities. First, the system uses a simple point-and-click camera-based interface. This feature addresses the complexity of conventional US and produces consistent data even when operated by a novice user6. Second, the system allows the use of water rather than traditional US gel as an acoustic medium. Previously, the use of US gel allowed the formation of bubbles that interfered with quality image acquisition. Also, the US gel is messy and provides challenges for clean-up. Further, the water is warmed by the heat lamp and helps maintain the animal&#39;s body temperature. Third, the robotic US is faster, so artifacts from respiratory motion are not problematic. The increased imaging speed allows for the practical completion of high throughput data collection. Fourth, the robotic US obtains 3D images, and therefore makes 3D reconstruction of objects simple (Figure 4). MRI and other modalities are expensive, time-consuming, and not always available. Importantly, the robotic US system fits on a table or bench and is more cost-efficient. Finally, prior work demonstrated that robotic US could provide comparable measurement data to more expensive modalities, such as MRI3.
While the image quality and resolution of the robotic US system described in this work were adequate for the proposed application (Figure 7), there are several ways that image quality can be improved in the future. For instance, utilizing higher frequency transducers (e.g., 50-70 MHz) would result in higher resolution images with better feature definition. While utilizing higher frequencies would result in a poorer depth of penetration, the images should be sufficient for in vivo imaging of superficial organs, such as the mouse kidney. As with other imaging modalities, contrast agents may be used to enhance specific features. For US, this typically means using something highly reflective of sound waves. Intravascular microbubbles in which lipids surround very small gas bubbles are one such agent. The micron-sized gas bubbles are highly reflective and thus provide a second distinct signal that translates into a high-resolution of vasculature7. While this acoustic contrast technique may be quite useful, it can have several downsides. First, the microbubbles must be made fresh and only persist in vivo for 5-10 min. Second, in vivo intravascular injection typically requires tail veil catheterization for injection, and this can be technically challenging. Under certain circumstances and pulsing regimes, microbubble imaging can itself lead to renal vasculature damage8.
There are also some more general limitations of the particular US system used. First, only one linear array (centered at 18 MHz) is included in the robotic chassis, so switching to higher or lower frequency probes is not currently possible. This may impact the breadth of models (either larger or smaller) that can be evaluated with the system. Future iterations of the instrument should include multiple linear arrays to cover the full range of preclinical animal models. Second, the transducer angle relative to the animal subject cannot be controlled. Therefore, performing angle-dependent imaging techniques, such as Doppler, or achieving alternative in-plane views of certain organs (e.g., long axis view of kidney) requires repositioning of the animal and can be difficult to achieve. Additional degrees-of-freedom could be added to the robotic movement to ameliorate this challenge. Third, on occasion, we have observed reverberation artifacts arising from the acoustic membrane that separates the animal from the transducer that can obscure visualization of superficial features and boundaries. In these cases, using a gel standoff to elevate the animal away from the membrane can remedy the situation. Finally, temperature control via heat lamp is imprecise, and therefore close attention must be paid to the animal&#39;s core body temperature while imaging. More controlled heating mechanisms, such as an integrated heating pad, will likely improve homeostasis management and imaging throughput.
The use of robotic US may be applicable to various fields of research. This technology enables visualization of gross tissue structures, thus may be used to track tumor progression and potential therapies6,9 as well as renal morphology as presented here. The ability to segment the specific features of the images makes it an attractive tool for studying models of polycystic kidney disease (PKD)3. M-mode images allow for simple quantification of many important cardiac parameters enabling in vivo assessment of cardiac physiology. As the kidneys receive 20%-25% of the cardiac output10, understanding cardiac function during the longitudinal assessment of renal pathology is important. Through these US protocols, we have tried to illustrate that US imaging is not only practical for in vivo and longitudinal kidney studies but also that increasingly US tools enable both morphologic as well as physiologic assessment of mice in preclinical studies.

An erratum was issued for:&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/t/62682">Use of 3D Robotic Ultrasound for In Vivo Analysis of Mouse Kidneys</a>. An author name was updated, along with the Protocol and Representative Results.
The&#160;name of an&#160;author was updated from:
Ryan C. Gesser
to:
Ryan C. Gessner
Step 3.1 of the Protocol was updated from:
Turn on the US machine and the heat lamp above the US platforms. Turn on the appropriate anesthesia valves for the corresponding US bays that will be used.
to:
Turn on the US machine and the switch for the internal heating element for the US bays. Turn on the appropriate anesthesia valves for the corresponding US bays that will be used.
Figue 1 in the Representative Results was updated from:
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62682/62682fig1v2.jpg" /> 
Figure 1: Ultrasound system and mouse placement. (A) Diagram of ultrasound system and location of transducers. (B) View of mice in supine position on ultrasound platform with heat lamp. (C) Example of region of interest (ROI)s in place for area of interest (kidneys) with animal IDs. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62682/62682fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
to:
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62682/62682fig1v3.jpg" /> 
Figure 1: Ultrasound system and mouse placement. (A) Diagram of ultrasound system and location of transducers. (B) View of mice in supine position on ultrasound platform. (C) Example of region of interest (ROI)s in place for area of interest (kidneys) with animal IDs. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62682/62682fig1v3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

An erratum was issued for:&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/t/62682">Use of 3D Robotic Ultrasound for In Vivo Analysis of Mouse Kidneys</a>. An author name was updated, along with the Protocol and Representative Results.

Erratum: Use of 3D Robotic Ultrasound for In Vivo Analysis of Mouse Kidneys

The most common modalities for in vivo rodent imaging include positron emission tomography (PET), optical imaging (OI), computed tomography (CT), magnetic resonance imaging (MRI), and ultrasound (US). These techniques provide high-resolution in vivo images, allowing investigators to quantitatively assess and longitudinally follow disease models non-invasively1. While each imaging modality has limitations, they also provide invaluable tools for preclinical research.
Here, the study details a US system and presents the protocol for robotic and 3D rodent imaging. US waves are produced by a probe called a transducer, which is typically hand-held. Sound waves are reflected back as they interact with tissues, and the echoes are reconstructed into images2. The protocol described here will focus on kidney and cardiac imaging using a robotically controlled transducer and using software that allows rapid 3D reconstruction for quantitative assessment.
Robotic US is a fast, reliable, and non-invasive imaging modality that allows investigators to conduct high-throughput and longitudinal studies. Compared to hand-held US methods, the robotic US method is time-efficient, as up to three animals can be scanned in a matter of minutes. High throughput for kidney measurements suggests that up to 20 mice per hour may be imaged. The robotic transducers are located underneath the acoustic membranes and move independently of the animal with two degrees of freedom (Figure 1A). This allows novice users to obtain high-quality images, whereas hand-held US methods are more susceptible to user error. The coupled software allows efficient, real-time 3D kidney reconstruction. Previously, magnetic resonance imaging (MRI) has been a prevalent method for non-invasive imaging due to the excellent soft-tissue contrast, lack of radioactivity, and penetration depth. However, MRI often requires long acquisition times and is costly to perform. US has been evaluated as a reliable and more rapid alternative to MRI in assessing total kidney volume (TKV)3.

<table><tbody><tr><td>Electric Razor</td><td>Braintree Scientific, Inc</td><td>CLP-9868 14</td><td></td></tr><tr><td>C57bk6j</td><td>The Jackson laboratory</td><td></td><td>https://www.jax.org/</td></tr><tr><td>Cotton gauze pads</td><td>Fisher Scientific</td><td></td><td></td></tr><tr><td>Cotton tipped applicators</td><td>Fisher Scientific</td><td></td><td></td></tr><tr><td>Depilatory cream</td><td>N/a</td><td>N/a</td><td>This study used Nair</td></tr><tr><td>Heat lamp</td><td></td><td></td><td>Included with SonoVol Vega system</td></tr><tr><td>Robotic Ultrasound System</td><td>SonoVol Inc</td><td></td><td>SonoVol Vega system includes anesthesia system</td></tr><tr><td>SonoEQ Software</td><td>SonoVol</td><td></td><td>Included with SonoVol Vega system</td></tr><tr><td>TERRELL Isoflurane</td><td>Piramal Critical Care, Inc</td><td>NDC 66794-019-10</td><td></td></tr></tbody></table>

use of 3d robotic ultrasound for in vivo analysis of mouse kidneys

Common modalities for in vivo imaging of rodents include positron emission tomography (PET), computed tomography (CT), magnetic resonance imaging (MRI), and ultrasound (US). Each method has limitations and advantages, including availability, ease of use, cost, size, and the use of ionizing radiation or magnetic fields. This protocol describes the use of 3D robotic US for in vivo imaging of rodent kidneys and heart, subsequent data analysis, and possible research applications. Practical applications of robotic US are the quantification of total kidney volume (TKV), as well as the measurement of cysts, tumors, and vasculature. Although the resolution is not as high as other modalities, robotic US allows for more practical high throughput data collection. Furthermore, using US M-mode imaging, cardiac function may be quantified. Since the kidneys receive 20%-25% of the cardiac output, assessing cardiac function is critical to the understanding of kidney physiology and pathophysiology.

Results of kidney analysis 
Surface area and volume data are acquired from the segmentation of the kidneys. This information can be used to compare experimental and control models or track changes over time. The calipers tool is useful for quickly measuring abnormalities (i.e., cysts, tumors) and how they change in length over time. Figure 3 suggests that both the segmentation and caliper methods can be used to measure cyst volumes accurately. Figure 4 demonstrates a clear difference in total kidney volume (TKV) between age-matched control and experimental (Pkd1RC/RC) mice. 3D visualization of these volume renderings may be performed within the system, including rotations within 3D space (Figure&#160;5). These 3D-reconstructions are then used to calculate TKV (mm3; Figure 4) as well as individual large cyst volume.
Results of cardiac analysis 
Many useful parameters are acquired from the analysis of M Mode images. These data provide a good snapshot of the left ventricular (LV) cardiac function at that point in time. The data output includes LV internal diameter, LV posterior wall, LV anterior wall diameter, ejection fraction, fractional shortening, stroke volume, heart rate, cardiac output, LV volume, and LV mass. The success of cardiac analysis is dependent on accurate segmentation of the layers on the M Mode image. Most cardiovascular results are calculated by the peak systolic and diastolic phases of the posterior and anterior endocardial layers. The posterior epicardial layer appears bright white and follows a similar pattern to the posterior endocardial layer. The tracing for the posterior endocardial layer should be placed on the lowest contour. The anterior endocardial layer should be traced along the highest contour of that layer. The anterior epicardial layer appears linear at the bottom of the image due to the prone positioning of the animal (Figure 2D). Figure 6 shows an example of one study with no significant difference in cardiac output between experimental and control mice. As with renal imaging, 3D cardiac visualization is possible. Nevertheless, a 4D visualization of the cardiac cycle (Supplemental Figure 1) allows the investigator to visualize and pinpoint both morphologic and cycle-dynamic abnormalities in the assessed animal.
Morphology assessment 
For quick and inexpensive assessment, US can effectively monitor physiological parameters longitudinally. However, many studies wish to additionally determine finer morphologic characteristics, e.g., number and sizes of cysts, calcifications (kidney stones), vascularization, or degree of fibrosis. Figure 7 compares a normal mouse kidney to a cystic mouse kidney to a moderately calcified mouse kidney. By increasing the US center frequency (10 MHz with the linear array) to 35 MHz (wobbler amplifier), pictures of increasing detail may be obtained.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62682/62682fig1v3.jpg" /> 
Figure 1: Ultrasound system and mouse placement. (A) Diagram of ultrasound system and location of transducers. (B) View of mice in supine position on ultrasound platform. (C) Example of region of interest (ROI)s in place for area of interest (kidneys) with animal IDs. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62682/62682fig1v3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62682/62682fig02.jpg" /> 
Figure 2: Cardiac ultrasound imaging to obtain physiological parameters. (A) Use of the Heart Finder heatmap image to position the transducer in the left ventricle for M-Mode imaging. The transducer location in the left ventricle is indicated by the large green dot. (B) View of the transducer when placed correctly over papillary muscles (dotted box). (C) Example view of layers needed to measure cardiac parameters. (D) View of live M-Mode image with layers designated as in panel C. (Layers from top to bottom: posterior epicardial, posterior endocardial, anterior endocardial, and anterior epicardial.) (E) Example output of statistics generated from cardiac measurements. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62682/62682fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62682/62682fig03.jpg" /> 
Figure 3: Using segmentation and calipers tools to measure kidneys and cyst. (A) Example segmentations (axial view) of both kidneys (blue and orange shading) and a large cyst (yellow) with volumes listed below. Non-segmented views are shown underneath so that the unobscured US may be viewed. (B) Example use of calipers to measure the same cyst (sagittal view) from Figure 3A with measurements below. The volume was calculated using the formula for an ellipse (volume = (4/3)&#960; x a x b x c, where a, b, c are relative x, y, z, respectively). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62682/62682fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62682/62682fig04.jpg" /> 
Figure 4: TKV distributions of WT and cystic mouse kidneys. Representation of TKVs for wild-type (WT) (C57BL/6J) and diseased (Pkd1RC/RC) mice. n = 22 (WT) n = 9 (Pkd1RC/RC); Results of two-tailed t-test: p &#60; 0.0001. Box shows 25-75th percentile values and whiskers show 1.5 times interquartile range. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62682/62682fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62682/62682fig05.jpg" /> 
Figure 5: Animated 3D reconstruction of segmented kidneys and cyst. Using the software, the 3D projections of the kidneys and cyst may be rotated or rocked in the 3D space (blue = left kidney; yellow = large cyst; orange = right kidney). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62682/Figure05.pptx" target="_blank">Please click here to download this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/62682/62682fig06.jpg" /> 
Figure 6: Cardiac physiological parameters from US measurements. Representation of cardiac output (mL/min) for WT and diseased (Pkd1RC/RC) mice. n = 22 (WT) n = 9 (Pkd1RC/RC). The lower tabulated data show that there is no significant difference for these two groups in ejection fraction, stroke volume, heart rate (HR), or cardiac output (CO). Results of two-tailed t-test: p &#62; 0.05. Box shows 25-75th percentile values. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62682/62682fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/62682/62682fig07.jpg" /> 
Figure 7: Comparison of US sagittal sections of normal and two pathologies. (A) Wild-type (C57BL/6J&#160;strain) kidney (TKV = 143.202 mm3). (B) Cystic kidney with increased TKV (Pkd1RC/RC mouse) (TKV = 333.158 mm3). Cysts are indicated by yellow arrows. (C) Kidney with vascular calcifications (Model = Low-Density Lipoprotein Receptor Deficient, Apolipoprotein B100-only mouse fed a Western diet for 12 months5) (TKV = 127.376 mm3). Renal stones are indicated by green arrows. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62682/62682fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Figure 1: 4D cardiac cycle movie from US measurements.&#160;Using the software, a representation of the beating heart is captured in 3D US and projected through the cardiac cycle. The green arrow indicates the aortic valve. (Model = Low-Density Lipoprotein Receptor Deficient, Apolipoprotein B100-only mouse, fed a Western diet for 12 months5). This model generates vascular calcifications enabling easier visualization of the heart and valves due to the greater acoustic reflectivity of the calcifications in US. Similar 4D reconstructions are possible with WT mice; however, the captured acoustic contrast will not be as high. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62682/SupplFigure01.pptx" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Use of 3D Robotic Ultrasound for In Vivo Analysis of Mouse Kidneys at JoVE.com

Use of 3D Robotic Ultrasound for In Vivo Analysis of Mouse Kidneys

This protocol demonstrates robotic ultrasound (US) as a practical, cost-effective, and quick alternative to traditional non-invasive image modalities.

Use of 3D Robotic Ultrasound for In Vivo Analysis of Mouse Kidneys

Common modalities for in vivo imaging of rodents include positron emission tomography (PET), computed tomography (CT), magnetic resonance imaging (MRI), ...

use-of-3d-robotic-ultrasound-for-in-vivo-analysis-of-mouse-kidneys

Nanyang Technological University

Research

JoVE Journal

Medicine

3.4K Views.  Mayo Clinic. Met dit protocol kunnen beginnende gebruikers hoogwaardige nier-, hart- en levergegevens verkrijgen. De geïntegreerde robotische echografie is ook belangrijk, omdat het niet nodig is om dieren of de akoestische sondes te verplaatsen. De belangrijkste voordelen van deze techniek zijn zowel snelheid als gebruiksgemak van het instrument.Men kan gemakkelijk beelden van hoge kwaliteit verkrijgen van 20 tot 30 muizen in een uur. We zouden verwachten dat een beginnende gebruiker moeite heeft...

Video: Use of 3D Robotic Ultrasound for In Vivo Analysis of Mouse Kidneys

Murine Echocardiography and Ultrasound Imaging

Rodent models of cardiac pathophysiology represent a valuable research tool to investigate mechanism of disease as well as test new therapeutics.1 Echocardiography provides a powerful, non-invasive tool to serially assess cardiac morphometry and function in a living animal.2 However, using this technique on mice poses unique challenges owing to the small size and rapid heart rate of these animals.3 Until recently, few ultrasound systems were capable of performing quality echocardiography on mice, and those generally lacked the image resolution and frame rate necessary to obtain truly quantitative measurements. Newly released systems such as the VisualSonics Vevo2100 provide new tools for researchers to carefully and non-invasively investigate cardiac function in mice. This system generates high resolution images and provides analysis capabilities similar to those used with human patients. Although color Doppler has been available for over 30 years in humans, this valuable technology has only recently been possible in rodent ultrasound.4,5 Color Doppler has broad applications for echocardiography, including the ability to quickly assess flow directionality in vessels and through valves, and to rapidly identify valve regurgitation. Strain analysis is a critical advance that is utilized to quantitatively measure regional myocardial function.6 This technique has the potential to detect changes in pathology, or resolution of pathology, earlier than conventional techniques. Coupled with the addition of three-dimensional image reconstruction, volumetric assessment of whole-organs is possible, including visualization and assessment of cardiac and vascular structures. Murine-compatible contrast imaging can also allow for volumetric measurements and tissue perfusion assessment.

This video demonstrates use of a rail-mounted high-frequency ultrasound probe to perform echocardiography on an anesthetized mouse. The methods describe both conventional two-dimensional and M-mode measurements of cardiac function in addition to newer, more powerful tools such as color Doppler, strain analysis, as well as general and targeted contrast imaging.

Rodent models of cardiac pathophysiology represent a valuable research tool to investigate mechanism of disease as well as test new therapeutics.1  ...

A Novel Use of Three-dimensional High-frequency Ultrasonography for Early Pregnancy Characterization in the Mouse

High-frequency ultrasonography (HFUS) is a common method to non-invasively monitor the real-time development of the human fetus in utero. The mouse is routinely used as an in vivo model to study embryo implantation and pregnancy progression. Unfortunately, such murine studies require pregnancy interruption to enable follow-up phenotypic analysis. To address this issue, we used three-dimensional (3-D) reconstruction of HFUS imaging data for early detection and characterization of murine embryo implantation sites and their individual developmental progression in utero. Combining HFUS imaging with 3-D reconstruction and modeling, we were able to accurately quantify embryo implantation site number as well as monitor developmental progression in pregnant C57BL6J/129S mice from 5.5 days post coitus (d.p.c.) through to 9.5 d.p.c. with the use of a transducer. Measurements included: number, location, and volume of implantation sites as well as inter-implantation site spacing; embryo viability was assessed by cardiac activity monitoring. In the immediate post-implantation period (5.5 to 8.5 d.p.c.), 3-D reconstruction of the gravid uterus in both mesh and solid overlay format enabled visual representation of the developing pregnancies within each uterine horn. As genetically engineered mice continue to be used to characterize female reproductive phenotypes derived from uterine dysfunction, this method offers a new approach to detect, quantify, and characterize early implantation events in vivo. This novel use of 3-D HFUS imaging demonstrates the ability to successfully detect, visualize, and characterize embryo-implantation sites during early murine pregnancy in a non-invasive manner. The technology offers a significant improvement over current methods, which rely on the interruption of pregnancies for gross tissue and histopathologic characterization. Here we use a video and text format to describe how to successfully perform ultrasounds of early murine pregnancy to generate reliable and reproducible data with reconstruction of the uterine form in mesh and solid 3-D images.

Mice are widely used to study gestational biology. However, pregnancy termination is required for such studies which precludes longitudinal investigations and necessitates the use of large numbers of animals. Therefore, we describe a non-invasive technique of high-frequency ultrasonography for early detection and monitoring of post-implantation events in the pregnant mouse.

High-frequency ultrasonography (HFUS) is a common method to non-invasively monitor the real-time development of the human fetus in utero. The mouse is ...

Developmental Biology

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 ...

Cancer Research

Early Detection of Drug-Induced Renal Hemodynamic Dysfunction Using Sonographic Technology in Rats

The kidney normally functions to maintain hemodynamic homeostasis and is a major site of damage caused by drug toxicity. Drug-induced nephrotoxicity is estimated to contribute to 19- 25% of all clinical cases of acute kidney injury (AKI) in critically ill patients. AKI detection has historically relied on metrics such as serum creatinine (sCr) or blood urea nitrogen (BUN) which are demonstrably inadequate in full assessment of nephrotoxicity in the early phase of renal dysfunction. Currently, there is no robust diagnostic method to accurately detect hemodynamic alteration in the early phase of AKI while such alterations might actually precede the rise in serum biomarker levels. Such early detection can help clinicians make an accurate diagnosis and help in in decision making for therapeutic strategy. Rats were treated with Cisplatin to induce AKI. Nephrotoxicity was assessed for six days using high-frequency sonography, sCr measurement and upon histopathology of kidney. Hemodynamic evaluation using 2D and Color-Doppler images were used to serially study nephrotoxicity in rats, using the sonography. Our data showed successful drug-induced kidney injury in adult rats by histological examination. Color-Doppler based sonographic assessment of AKI indicated that resistive-index (RI) and pulsatile-index (PI) were increased in the treatment group; and peak-systolic velocity (mm/s), end-diastolic velocity (mm/s) and velocity-time integral (VTI, mm) were decreased in renal arteries in the same group. Importantly, these hemodynamic changes evaluated by sonography preceded the rise of sCr levels. Sonography-based indices such as RI or PI can thus be useful predictive markers of declining renal function in rodents. From our sonography-based observations in the kidneys of rats that underwent AKI, we showed that these noninvasive hemodynamic measurements may consider as an accurate, sensitive and robust method in detecting early stage kidney dysfunction. This study also underscores the importance of ethical issues associated with animal use in research.

Early stage hemodynamic dysfunction is critical to the development of kidney disease. Yet, detection methodologies are limited. Recent advances in sonography provide a noninvasive, accurate option for early detection of kidney injury. This study outlines a step-by-step, sonographic methodology for detecting kidney dysfunction using a drug-induced nephrotoxicity rat model.

The kidney normally functions to maintain hemodynamic homeostasis and is a major site of damage caused by drug toxicity. Drug-induced nephrotoxicity is ...

Use of Ultra-high Field MRI in Small Rodent Models of Polycystic Kidney Disease for In Vivo Phenotyping and Drug Monitoring

Several in vivo pre-clinical studies in Polycystic Kidney Disease (PKD) utilize orthologous rodent models to identify and study the genetic and molecular mechanisms responsible for the disease, and are very convenient for rapid drug screening and testing of promising therapies. A limiting factor in these studies is often the lack of efficient non-invasive methods for sequentially analyzing the anatomical and functional changes in the kidney. Magnetic resonance imaging (MRI) is the current gold standard imaging technique to follow autosomal dominant polycystic kidney disease (ADPKD) patients, providing excellent soft tissue contrast and anatomic detail and allowing Total Kidney Volume (TKV) measurements.A major advantage of MRI in rodent models of PKD is the possibility for in vivo imaging allowing for longitudinal studies that use the same animal and therefore reducing the total number of animals required. In this manuscript, we will focus on using Ultra-high field (UHF) MRI to non-invasively acquire in vivo images of rodent models for PKD. The main goal of this work is to introduce the use of MRI as a tool for in vivo phenotypical characterization and drug monitoring in rodent models for PKD.

Use of Ultra-high Field MRI in Small Rodent Models of Polycystic Kidney Disease for In Vivo Phenotyping and Drug Monitoring

The use of ultra-high field MRI as a non-invasive way to obtain phenotypic information of rodent models for polycystic kidney disease and to monitor interventions is described. Compared with the traditional histological approach, MRI images can be acquired in vivo, allowing for longitudinal follow-up.

Several in vivo pre-clinical studies in Polycystic Kidney Disease (PKD) utilize orthologous rodent models to identify and study the genetic and molecular ...

Using 2-Photon Microscopy to Quantify the Effects of Chronic Unilateral Ureteral Obstruction on Glomerular Processes

Applying novel microscopy methods to suitable animal disease models to explore the dynamic physiology of the kidney remains a challenge. Rats with surface glomeruli provide a unique opportunity to investigate physiological and pathophysiological processes using intravital 2-photon microscopy. Quantification of glomerular capillary blood flow and vasoconstriction and dilatation in response to drugs, permeability, and inflammation are just some of the processes that can be studied. In addition, transgenic rats, i.e., podocytes labeled with fluorescent dyes and other molecular biomarker approaches, provide increased resolution to directly monitor and quantify protein-protein interactions and the effects of specific molecular alterations.
In mice, which lack surface glomeruli after four weeks of age, unilateral ureteral obstruction (UUO) for several weeks has been used to induce surface glomeruli. As this induction model does not allow for baseline studies, we quantified the effects of UUO on glomerular processes in the UUO model in Munich Wistar Fr&#246;mter (MWF) rats, which have surface glomeruli under physiologic conditions. The UUO model for five weeks or more induced significant alterations to gross renal morphology, the peritubular and glomerular microvasculature, as well as the structure and function of tubular epithelia. Glomerular and peritubular red blood cell (RBC) flow decreased significantly (p &#60; 0.01), probably due to the significant increase in the adherence of white blood cells (WBCs) within glomerular and peritubular capillaries. The glomerular sieving coefficient of albumin increased from 0.015 &#177; 0.002 in untreated MWFs to 0.045 &#177; 0.05 in 5-week-old UUO MWF rats. Twelve weeks of UUO resulted in further increases in surface glomerular density and glomerular sieving coefficient (GSC) for albumin. Fluorescent albumin filtered across the glomeruli was not reabsorbed by the proximal tubules. These data suggest that using UUO to induce surface glomeruli limits the ability to study and interpret normal glomerular processes and disease alterations.

Here, we present a protocol using 2-photon microscopy in Munich Wistar Fromter rats with surface glomeruli to quantifythe effects of prolonged ureteral obstruction on glomerular dynamics and function.

Applying novel microscopy methods to suitable animal disease models to explore the dynamic physiology of the kidney remains a challenge. Rats with surface ...

Whole-Kidney Three-Dimensional Staining with CUBIC

Although conventional pathology provided numerous information about kidney microstructure, it was difficult to know the precise structure of blood vessels, proximal tubules, collecting ducts, glomeruli, and sympathetic nerves in the kidney due to the lack of three-dimensional (3D) information. Optical clearing is a good strategy to overcome this big hurdle. Multiple cells in a whole organ can be analyzed at single-cell resolution by combining tissue clearing and 3D imaging technique. However, cell labeling methods for whole-organ imaging remain underdeveloped. In particular, whole-mount organ staining is challenging because of the difficulty in antibody penetration. The present protocol developed a whole-mount mouse kidney staining for 3D imaging with the CUBIC (Clear, Unobstructed Brain/Body Imaging Cocktails and Computational analysis) tissue clearing method. The protocol has enabled visualizing renal sympathetic denervation after ischemia-reperfusion injury and glomerulomegaly in the early stage of diabetic kidney disease from a comprehensive viewpoint. Thus, this technique can lead to new discoveries in kidney research by providing a macroscopic perspective.

The present protocol describes a tissue clearing method and whole-mount immunofluorescent staining for three-dimensional (3D) kidney imaging. This technique can offer macroscopic perspectives in kidney pathology, leading to new biological discoveries.

Although conventional pathology provided numerous information about kidney microstructure, it was difficult to know the precise structure of blood ...

Drug Treatment by Central Venous Catheter in a Mouse Model of Angiotensin II Induced Abdominal Aortic Aneurysm and Monitoring by 3D Ultrasound

Since pharmaceutical treatment options are lacking in the clinical management of abdominal aortic aneurysm (AAA), animal models, in particular mouse models, are applied to advance the understanding of the disease pathogenesis and to identify potential therapeutic targets. Testing novel drug candidates to block AAA growth in these models generally requires repeated drug administration during the time course of the experiment. Here, we describe a compiled protocol for AAA induction, insertion of an intravenous catheter to facilitate prolonged therapy, and serial AAA monitoring by 3D ultrasound. Aneurysms are induced in apolipoprotein E (ApoE) deficient mice by angiotensin II release over 28 days from osmotic mini-pumps implanted subcutaneously into the mouse back. Subsequently, the surgical procedure for external jugular vein catheterization is conducted to allow for daily intravenous drug treatment or repeated blood sampling via a subcutaneous vascular access button. Despite the two dorsal implants, the monitoring of AAA development is readily facilitated by sequential semi-automated 3D ultrasound analysis, which yields comprehensive information on the expansion of aortic diameter and volume and on aneurysm morphology, as illustrated by experimental examples.

This protocol describes the consecutive implantation of an osmotic pump to induce abdominal aortic aneurysm by angiotensin II release in apolipoprotein E (ApoE) deficient mice and of a vascular access port with a jugular vein catheter for repeated drug treatment. Monitoring of aneurysm development by 3D ultrasound is effectively conducted despite dorsal implants.

Since pharmaceutical treatment options are lacking in the clinical management of abdominal aortic aneurysm (AAA), animal models, in particular mouse ...

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 ...

Fluorescent Labeling of Glomeruli in Cleared Mouse Kidneys for Microscopy

An Efficient and Fast Method for Labeling and Analyzing Mouse Glomeruli

The glomeruli are fundamental units in the kidney; hence, studying the glomeruli is pivotal for understanding renal function and pathology. Biological imaging provides intuitive information; thus, it is of great significance to label and observe the glomeruli. However, the glomeruli observation methods currently in use require complicated operations, and the results may lose label details or three-dimensional (3D) information. The clear, unobstructed brain imaging cocktails and computational analysis (CUBIC) tissue clearing technology has been widely used in renal research, allowing for more accurate detection and deeper detection depth. We found that mouse glomeruli can be rapidly and effectively labeled by tail vein injection of medium molecular weight FITC-Dextran followed by the CUBIC clearing method. The cleared mouse kidney could be scanned by a light-sheet microscope (or a confocal microscope when sliced) to obtain three-dimensional image stacks of all the glomeruli in the entire kidney. Processed with appropriate software, the glomeruli signals could be easily digitized and further analyzed to measure the number, volume, and frequency of the glomeruli.

Author Spotlight: Aiding Research in Kidney Biology by Labeling Glomeruli in Cleared Tissues 

This study presents an easy-to-use, complete, and simple set of methods to label and analyze glomeruli from CUBIC-cleared mouse kidneys. Data such as glomerulus number and volume can be obtained easily and reliably using fluorescein isothiocyanate (FITC)-Dextran, light sheet fluorescence microscopy (LSFM), or common confocal microscopy and software such as Imaris.

The glomeruli are fundamental units in the kidney; hence, studying the glomeruli is pivotal for understanding renal function and pathology. Biological ...

Use of 3D Robotic Ultrasound for In Vivo Analysis of Mouse Kidneys

Samenvatting

Meer video's verkennen