This work was supported by R01HL124155 (CPH) and funding from the Research Institute at the University of Missouri to CPH.

The protocol for handling of mice and ultrasound imaging was approved by the University of Missouri Institutional Animal Care and Use Committee (animal protocol number 8799) and was conducted according to AAALAC International.
1. Equipment setup and preparation of mice
<ol>
	<li>Equipment setup
	<ol>
		<li>Turn on the ultrasound instrument, ultrasonic gel warmer and the heating pad.</li>
		<li>Open the ultrasound program and enter the study name and descriptive information for each mouse.</li>
		<li>Select the application as General Imaging.</li>
		<li>Choose the appropriate transducer for abdominal imaging (Figure 1B,C). In this experiment, MS400 transducer is used.</li>
		<li>Ensure anesthesia isoflurane and oxygen levels are adequate for each experimental session.</li>
		<li>Clean the ultrasound animal imaging platform.</li>
	</ol>
	</li>
	<li>Mouse preparation
	<ol>
		<li>Place the mouse cage on top of a heating pad (36.5 to 38.5 &#176;C).</li>
		<li>Gently hold the mouse by its tail base and place in the oxygen-filled isoflurane chamber.</li>
		<li>Direct the isoflurane and oxygen flow to the induction chamber.</li>
		<li>Turn on the isoflurane vaporizer and set the isoflurane level to 1-2% vol/vol. Turn on the oxygen tank pressure to 1-2 L/min.</li>
		<li>After ~2 min, confirm the adequate depth of anesthesia by the absence of withdrawal reflexes upon pinching the foot pad of the mouse.</li>
		<li>Next, turn off the induction chamber supply branch and turn on the branch directed to the anesthesia nose cone.</li>
		<li>Transfer the mouse from the induction chamber to the ultrasound imaging stage and position the anesthesia cone over the nose of the animal.</li>
		<li>Tilt the animal imaging platform around 10&#176; to the lower right corner for optimal scanning (Figure 1B).</li>
		<li>Put one drop of sterile ophthalmic solution in both eyes of mice to prevent drying under anesthesia.</li>
		<li>Position the mouse in the supine position with its nose inserted into the anesthesia cone.</li>
		<li>Apply the electrode gel to all four paws using a cotton swab and tape the paws to the copper leads on the animal imaging platform for electrocardiogram readings (Figure 1C).</li>
		<li>Use clippers to shave hair at the imaging site and then apply depilatory cream to remove remaining fur. Leave for less than 1 min.</li>
		<li>Gently wipe off the cream and hair with a damp paper towel.</li>
		<li>Monitor the breathing and ensure that heart rate is maintained between 450-550 beats/min. If below this level, reduce the isoflurane flow and wait until the heart rate recovers.</li>
		<li>Apply prewarmed ultrasonic gel (37 &#176;C) to the prepared skin site and attach the transducer to its holder and lower down until it touches the gel (Figure 1C).</li>
	</ol>
	</li>
</ol>
2. Ultrasound imaging of the abdominal aorta
<ol>
	<li>Position the transducer horizontally (i.e., perpendicular to the midline of the mouse).</li>
	<li>Smooth the ultrasonic gel and remove bubbles using the wood stick of a cotton swab.</li>
	<li>Lower the transducer and place 0.5 - 1 cm below the diaphragm after touching the gel. Now start to observe the images.</li>
	<li>Visualize the abdominal aorta in the short axis view (Figure 1C). 
	NOTE: B-mode is the default and most effective mode to anatomically locate the aorta and position the transducer. The abdominal aorta is identified by the presence of pulsatile flow using color Doppler and power Doppler modes in the short axis (i.e., the circumferential cross-section of the aorta). Adjust the micromanipulators on the animal stage and the transducer to bring the cross section of the aorta to the center of the image.</li>
	<li>Gently rotate the transducer 90&#176; clockwise, and slowly adjust the x-axis micromanipulator knob to visualize the aorta in long axis view (longitudinal section of the aorta). 
	NOTE: In many cases, gastro-intestinal gases may interfere with the image, or the aorta may not be at the optimal angle to allow a clear long axis view. Adjust the angle of the transducer slowly and horizontally until an acceptable long axis view is obtained. If problems persist, elevate the transducer, check for air bubbles under the transducer, slightly adjust the tilting angle of the animal stage, reapply gels, and repeat all the steps again.</li>
	<li>Set the focus zone and depth at the region of the aorta using the Focus Zone and Focus Depth toggles, respectively. Adjust the time gain compensation slider manually to darken the lumen of the aorta to achieve an optimal contrast of the aorta wall.</li>
	<li>Adjust the y-axis manipulator to visualize the branching points of the superior mesenteric and the right renal arteries. Use the right renal artery as a landmark to capture image of the suprarenal aorta (Figure 2A).</li>
	<li>Record at least 100 frames of B-mode images on the suprarenal aorta.</li>
	<li>Press cinestore to save the B-mode images.</li>
	<li>Press M-mode button on the instrument keyboard to enable M-mode recording. Roll the cursor ball to bring the yellow indicator line to a normal aorta sections with clear vessel wall image, or to the sections where maximal diameter of aneurysm is observed.</li>
	<li>Press the SV/gate toggle and adjust the cursor ball to ensure that vessel walls are included in the measurement bracket. Press update to record M-mode measurements and press cinestore to capture (Figure 2A,B). 
	NOTE: Maximal diameter of the aneurysm may not be in the same imaging plane as the optimal long axis view of the aorta. Adjust the x-axis manipulator knob slightly for each M-mode measurement to ensure that the MILD of each section is captured.</li>
	<li>To obtain ECG-gated Kilohertz Visualization (EKV) images, press the B-mode button to go back to B-mode recording. 
	NOTE: If the images are not sharp, adjust the x-axis manipulator to achieve the sharpest image of upper wall of the lumen over a section length (i.e., &#62; 6 mm).</li>
	<li>Press Physio Settings button on the keyboard and select Respiration Gating. Adjust the gating Delay and Window manually to record the data only during the flatter parts of the respiration wave. The recording sections will be shown as colored blocks on the tracing of the respiration wave. 
	NOTE: Without the adjustment of the respiration gating, the EKV images will be blurred due to the normal movement of animal during breathing.</li>
	<li>Press EKV button to enable the EKV mode. In the appropriate menu, select Standard Resolution and frame rate 3000 or higher. Select proceed to record EKV images. Press cinestore to save the images. Use EKV mode image to obtain measurements of pulse propagation velocity (PPV), distensibility and radial strain. 
	NOTE: EKV recording may fail if there are abnormal fluctuations in respiration, animal is respiring too rapidly, or frame rates settings are too high. In those case, set the frame rate lower and wait for the animal respiration to stabilize. Setting the frame rate at 3000 is usually appropriate for both mice and rats.</li>
</ol>
3. Post-imaging steps
<ol>
	<li>Gently wipe the ultrasonic gel from the abdominal area of the mouse with a paper towel moistened with warm water.</li>
	<li>Place the mouse back in its home cage on a heating pad.</li>
	<li>Turn off the isoflurane machine, clean the animal imaging platform and transducer with damp wipes.</li>
	<li>Transfer the image data collected during the ultrasound scan to the hard drive.</li>
	<li>Turn off the ultrasound instrument.</li>
	<li>After the mouse recovers from anesthesia and is alert, remove the heating pad and return the cage to the animal housing rack.</li>
</ol>
4. Analysis of abdominal aortic images
<ol>
	<li>Analysis of M-mode images to measure MILD
	<ol>
		<li>Open the ultrasound program and enter the study name and descriptive information for each mouse.</li>
		<li>Open the ultrasound data in the analysis software and open the M-mode image and pause the heartbeat.</li>
		<li>Click on Measurements.</li>
		<li>Select the vascular package from the drop-down options. Click on Depth and draw a line across the aortic lumen extending from inner wall to wall (Figure 2C,D). 
		NOTE: For consistency, the measurements should be taken at the systolic phase of the cardiac cycle when the aorta is maximally expanded. Draw three lines across three different heartbeats to obtain accurate and average measurements of MILD. In AAA, the measurements are taken at the maximal dilatation of the aorta. It is also advisable to fast the animals 4-6 h prior to collecting images to avoid interference from bowel motility and ensure image clarity.</li>
	</ol>
	</li>
	<li>Analysis for pulse propagation velocity (PPV)
	<ol>
		<li>Open the EKV image and pause the heartbeat.</li>
		<li>Open a new window on the analysis software (e.g., Vevo Vac) by clicking on the name icon.</li>
		<li>Click on the PPV option (arrow in Figure 3D). A small window will further appear with the image of the aorta.</li>
		<li>Draw a rectangular box by clicking on the upper vessel wall and dragging the pointer for about 4 mm covering both the walls of the suprarenal aorta. 
		NOTE: Keep the length of the box consistent (~4 mm) for all the images. The user can adjust the rectangular box by rotating to align the box and selecting the line then dragging to a new position on the vessel being analyzed to obtain the most appropriate and clear inflection of the pulse wave. The vertical lines of data from the rectangle will be displayed and identified as the Left (top image) and Right (bottom image) on the ROI. For a better visualization of the inflection of the pulse wave, it is sometimes useful to the draw box only on the upper wall as shown in Figure 3. The software will automatically calculate the PPV (m/s). However, it&#39;s always better to manually adjust the purple lines to set the exact inflection point on the pulse waves and PPV will change accordingly.</li>
		<li>Finally, select the Accept command to save the PPV values. Export the figures and the data to the data storage drive.</li>
	</ol>
	</li>
	<li>Analysis for distensibility and radial strain
	<ol>
		<li>Open the EKV image and pause the heartbeat.</li>
		<li>Click on the software icon. The software will open a new window.</li>
		<li>Click on the trace new ROI and draw a rectangular box on the both walls of the vessel. The software will automatically trace the upper and lower walls of the vessel. The user can adjust the trace to align on the wall by clicking on green points (Figure 4A,B).</li>
		<li>Now Accept the trace. The software will calculate the distensibility (1/Mpa) in the selected ROI.</li>
		<li>For the radial strain measurement, select the appropriate strain option from the menu bars on the top left. The images for radial strain and tangential strain will open.</li>
		<li>Obtain the value for radial strain (%) by moving the cursor on the peak of the curve. Export the data as images or in video format (Figure 4A,B).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Ultrasound imaging provides a powerful technique for determining functional properties of the aorta through measurements of PPV, distensibility and radial strain. These measurements are particularly instructive for studying mouse models of AAA and the in vivo approach allows for collection of longitudinal data that is potentially important to understanding temporal development of the aortic pathology. Specifically, measurements of in vivo aortic stiffness are determined locally in the abdominal aorta by PPV, distensibili...

An abdominal aortic aneurysm (AAA) represents a significant cardiovascular disease characterized by a permanent localized dilation of the aorta exceeding the original vessel diameter by 1.5 times1. AAA ranks among the top 13 causes of mortality in the United States2. The progression of AAA is attributed to the degeneration of the aortic wall and elastin fragmentation, ultimately leading to aortic rupture. These changes in the aortic wall may occur without a significant increase in the maximal intraluminal diameter (MILD), thus suggesting that MILD alone is not sufficient to predict the severity of the disease3. Therefore, additional factors need to be identified to detect initial changes in the aortic wall, which may guide early treatment options. The overall goal of this protocol is to provide a practical guide for assessing aortic functional properties using ultrasound imaging as characterized by measurements of pulse propagation velocity (PPV), distensibility and radial strain.
A well characterized experimental model to study AAA, first described by Daugherty and colleagues, involves subcutaneous infusion of angiotensin II (AngII) via osmotic pumps in Apoe-/- mice4. Precise measurement of MILD using ultrasound imaging has been instrumental in characterizing AAA in this mouse model5. Although histological changes during the development of AAA have been extensively studied, changes in the functional properties of the vessel wall such as aortic stiffness have not been well characterized. This protocol emphasizes the use of high-frequency ultrasound in combination with the sophisticated analyses as powerful tools for studying the temporal progression of AAA. Specifically, these approaches allow us to assess the functional properties of the vessel wall as measured by PPV, distensibility and radial strain.
Recent clinical studies in human subjects with AAA, as well as in the murine elastase-induced AAA model, suggest a positive correlation between aortic stiffness and aortic diameter6,7. PPV, an indicator of aortic stiffness, is accepted as an excellent measurement for quantifying changes in stiffness in vessel wall6,8. PPV is calculated by measuring the transit time of the pulse waveform at two sites along the vasculature, thus providing a regional assessment of aortic stiffness. We have recently demonstrated that increased aortic stiffness as measured by PPV, and at the cellular level as determined using atomic force microscopy, positively correlates with aneurysm development9. Further, the literature suggests that aortic stiffness may precede aneurysmal dilation and thus may provide useful information about regional intrinsic properties of the vessel wall during development of AAA10. Similarly, distensibility and strain measurements are the quantification tools to measure earlier changes of arterial fitness. Healthy arteries are flexible and elastic, whereas with increased stiffness and less elasticity, distensibility and strain is decreased. Here, we provide a practical guide and step by step protocol for the use of a high-frequency ultrasound system to measure MILD, PPV, distensibility and radial strain in mice. The protocol provides technical approaches that should be used in conjunction with the basic information provided by manuals for specific ultrasound imaging instruments and the accompanying video tutorial. Importantly, in our hands the described imaging protocol provides reproducible and accurate data that appear valuable in the study of the development and progression of experimental AAA.
To further demonstrate the utility of ultrasound imaging, we provide example images and measurements taken from our own studies aimed at using pharmacological approaches for preventing experimental AAA11. Specifically, notch signaling has been proposed to be involved in multiple aspects of vascular development and inflammation12. Using gene haploinsufficiency and pharmacologic approaches, we have previously demonstrated that Notch inhibition reduces the development of AAA in mice by preventing infiltration of macrophages at the site of vascular injury13,14,15. For the current article, using the pharmacological approach for Notch inhibition we focus on the relationship between aortic stiffness and factors relating to AAA. These studies illustrate that Notch inhibition reduces aortic stiffness, which is a measure of AAA progression11.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Angiotensin II</td>
			<td>Sigma</td>
			<td>A9525</td>
			<td></td>
		</tr>
		<tr>
			<td>Apoe-/- mice</td>
			<td>The Jackon lab</td>
			<td><img src="/files/ftp_upload/60515/60515mateq.jpg"/></td>
			<td></td>
		</tr>
		<tr>
			<td>Clippers</td>
			<td>WAHL</td>
			<td>1854</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton swab</td>
			<td>Q-tips</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DAPT</td>
			<td>Sigma</td>
			<td>D5942</td>
			<td></td>
		</tr>
		<tr>
			<td>Depilatory cream</td>
			<td>Nair</td>
			<td>LL9038</td>
			<td></td>
		</tr>
		<tr>
			<td>Electrode cream</td>
			<td>Sigma</td>
			<td>17-05</td>
			<td></td>
		</tr>
		<tr>
			<td>Gel warmer</td>
			<td>Thermasonic (Parker)</td>
			<td>82-03 (LED)</td>
			<td></td>
		</tr>
		<tr>
			<td>Heating pad</td>
			<td>Stryker</td>
			<td>T/pump professional</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>VetOne</td>
			<td>Fluriso TM</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane vaporizer</td>
			<td>Visualsonics</td>
			<td>VS4244</td>
			<td></td>
		</tr>
		<tr>
			<td>Lubricating ophthalmic ointment</td>
			<td>Lacri-lube</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Osmotic pumps</td>
			<td>Alzet</td>
			<td>Model 2004</td>
			<td></td>
		</tr>
		<tr>
			<td>Oxygen tank</td>
			<td>Air gas</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tranducer</td>
			<td>Visualsonics</td>
			<td>MS-400 or MS550D</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasonic gel</td>
			<td>Parker</td>
			<td>Aquasonic clear</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasound Imaging System</td>
			<td>Visualsonics</td>
			<td>Vevo 2100</td>
			<td></td>
		</tr>
		<tr>
			<td>Vevo Vasc Software</td>
			<td>Visualsonics</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

measurement of pulse propagation velocity, distensibility and strain in an abdominal aortic aneurysm mouse model

An abdominal aortic aneurysm (AAA) is defined as a localized dilation of the abdominal aorta that exceeds the maximal intraluminal diameter (MILD) by 1.5 times of its original size. Clinical and experimental studies have shown that small aneurysms may rupture, while a subpopulation of large aneurysms may remain stable. Thus, in addition to the measurement of intraluminal diameter of the aorta, knowledge of structural traits of the vessel wall may provide important information to assess the stability of the AAA. Aortic stiffening has recently emerged as a reliable tool to determine early changes in the vascular wall. Pulse propagation velocity (PPV) along with the distensibility and radial strain are highly useful ultrasound-based methods relevant for assessing aortic stiffness. The primary purpose of this protocol is to provide a comprehensive technique for the use of ultrasound imaging system to acquire images and analyze the structural and functional properties of the aorta as determined by MILD, PPV, distensibility and radial strain.

Representative M-mode images of the normal and aneurysmal abdominal aorta from mice are shown in Figure 2A and Figure 2B, respectively. The suprarenal abdominal aorta is identified by its location next to right renal artery and the superior mesenteric artery (Figure 2A). Representative images used for the calculation of MILD, at three different heartbeats of the systolic cardiac cyc...

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Measurement of Pulse Propagation Velocity, Distensibility and Strain in an Abdominal Aortic Aneurysm Mouse Model

This manuscript describes a detailed protocol for using high frequency ultrasound imaging to measure luminal diameter, pulse propagation velocity, distensibility and radial strain on a mouse model of abdominal aortic aneurysm.

An abdominal aortic aneurysm (AAA) is defined as a localized dilation of the abdominal aorta that exceeds the maximal intraluminal diameter (MILD) by 1.5 ...

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JoVE Journal

Immunology and Infection

6.0K Views.  University of Missouri. This manuscript describes a detailed protocol for using high frequency ultrasound imaging to measure luminal diameter, pulse propagation velocity, distensibility and radial strain on a mouse model of abdominal aortic aneurysm.

Video: Measurement of Pulse Propagation Velocity, Distensibility and Strain in an Abdominal Aortic Aneurysm Mouse Model

Colon Ascendens Stent Peritonitis (CASP) - a Standardized Model for Polymicrobial Abdominal Sepsis

Sepsis remains a persistent problem on intensive care units all over the world. Understanding the complex mechanisms of sepsis is the precondition for establishing new therapeutic approaches in this field. Therefore, animal models are required that are able to closely mimic the human disease and also sufficiently deal with scientific questions. The Colon Ascendens Stent Peritonitis (CASP) is a highly standardized model for polymicrobial abdominal sepsis in rodents. In this model, a small stent is surgically inserted into the ascending colon of mice or rats leading to a continuous leakage of intestinal bacteria into the peritoneal cavity. The procedure results in peritonitis, systemic bacteraemia, organ infection by gut bacteria, and systemic but also local release of several pro- and anti-inflammatory cytokines. The lethality of CASP can be controlled by the diameter of the inserted stent. A variant of this model, the so-called CASP with intervention (CASPI), raises opportunity to remove the septic focus by a second operation according to common procedures in clinical practice. CASP is an easily learnable and highly reproducible model that closely mimics the clinical course of abdominal sepsis. It leads way to study on questions in several scientific fields e.g. immunology, infectiology, or surgery.

Colon Ascendens Stent Peritonitis CASP - a Standardized Model for Polymicrobial Abdominal Sepsis

The Colon Ascendens Stent Peritonitis (CASP) is a highly standardized model for polymicrobial abdominal sepsis in rodents. This article describes the surgical procedure of CASP. The CASP model and its variants allow the systematic investigation of various problems concerning the subject of sepsis.

Sepsis remains a persistent problem on intensive care units all over the world. Understanding the complex mechanisms of sepsis is the precondition for ...

Measurement of Tactile Allodynia in a Murine Model of Bacterial Prostatitis

Uropathogenic Escherichia coli (UPEC) are pathogens that play an important role in urinary tract infections and bacterial prostatitis1. We have recently shown that UPEC have an important role in the initiation of chronic pelvic pain2, a feature of Chronic prostatitis/Chronic pelvic pain syndrome (CP/CPPS)3,4. Infection of the prostate by clinically relevant UPEC can initiate and establish chronic pain through mechanisms that may involve tissue damage and the initiation of mechanisms of autoimmunity5.
A challenge to understanding the pathogenesis of UPEC in the prostate is the relative inaccessibility of the prostate gland to manipulation. We utilized a previously described intraurethral infection method6 to deliver a clinical strain of UPEC into male mice thereby establishing an ascending infection of the prostate. Here, we describe our protocols for standardizing the bacterial inoculum7 as well as the procedure for catheterizing anesthetized male mice for instillation of bacteria.
CP/CPPS is primarily characterized by the presence of tactile allodynia4. Behavior testing was based on the concept of cutaneous hyperalgesia resulting from referred visceral pain8-10. An irritable focus in visceral tissues reduces cutaneous pain thresholds allowing for an exaggerated response to normally non-painful stimuli (allodynia). Application of normal force to the skin result in abnormal responses that tend to increase with the intensity of the underlying visceral pain. We describe methodology in NOD/ShiLtJ mice that utilize von Frey fibers to quantify tactile allodynia over time in response to a single infection with UPEC bacteria.

Infection of the prostate may be a contributing factor in mediating pelvic pain in chronic prostatitis. We describe the procedure for preparation of standardized bacterial inoculum, instillation of bacteria into the urethra of male mice and methodology for measuring tactile allodynia in mice over time.

Uropathogenic Escherichia coli (UPEC) are pathogens that play an important role in urinary tract infections and bacterial prostatitis1. We have recently ...

Following in Real Time the Impact of Pneumococcal Virulence Factors in an Acute Mouse Pneumonia Model Using Bioluminescent Bacteria

Pneumonia is one of the major health care problems in developing and industrialized countries and is associated with considerable morbidity and mortality. Despite advances in knowledge of this illness, the availability of intensive care units (ICU), and the use of potent antimicrobial agents and effective vaccines, the mortality rates remain high1. Streptococcus pneumoniae is the leading pathogen of community-acquired pneumonia (CAP) and one of the most common causes of bacteremia in humans. This pathogen is equipped with an armamentarium of surface-exposed adhesins and virulence factors contributing to pneumonia and invasive pneumococcal disease (IPD). The assessment of the in vivo role of bacterial fitness or virulence factors is of utmost importance to unravel S. pneumoniae pathogenicity mechanisms. Murine models of pneumonia, bacteremia, and meningitis are being used to determine the impact of pneumococcal factors at different stages of the infection. Here we describe a protocol to monitor in real-time pneumococcal dissemination in mice after intranasal or intraperitoneal infections with bioluminescent bacteria. The results show the multiplication and dissemination of pneumococci in the lower respiratory tract and blood, which can be visualized and evaluated using an imaging system and the accompanying analysis software.

Streptococcus pneumoniae is the leading pathogen causing severe community-acquired pneumonia and responsible for over 2&#160;million deaths worldwide. The impact of bacterial factors implicated in fitness or virulence can be monitored in real-time in an acute mouse pneumonia or bacteremia model using bioluminescent bacteria.

Pneumonia is one of the major health care problems in developing and industrialized countries and is associated with considerable morbidity and mortality. ...

Influenza Virus Propagation in Embryonated Chicken Eggs

Influenza infection is associated with about 36,000 deaths and more than 200,000 hospitalizations every year in the United States. The continuous emergence of new influenza virus strains due to mutation and re-assortment complicates the control of the virus and necessitates the permanent development of novel drugs and vaccines. The laboratory-based study of influenza requires a reliable and cost-effective method for the propagation of the virus. Here, a comprehensive protocol is provided for influenza A virus propagation in fertile chicken eggs, which consistently yields high titer viral stocks. In brief, serum pathogen-free (SPF) fertilized chicken eggs are incubated at 37 &#176;C and 55-60% humidity for 10 &#8211; 11 days. Over this period, embryo development can be easily monitored using an egg candler. Virus inoculation is carried out by injection of virus stock into the allantoic cavity using a needle. After 2 days of incubation at 37 &#176;C, the eggs are chilled for at least 4 hr at 4 &#176;C. The eggshell above the air sac and the chorioallantoic membrane are then carefully opened, and the allantoic fluid containing the virus is harvested. The fluid is cleared from debris by centrifugation, aliquoted and transferred to -80 &#176;C for long-term storage. The large amount (5-10 ml of virus-containing fluid per egg) and high virus titer which is usually achieved with this protocol has made the usage of eggs for virus preparation our favorable method, in particular for in vitro studies which require large quantities of virus in which high dosages of the same virus stock are needed.

Fertile chicken eggs are widely used to produce large amounts of human influenza A virus as they provide a convenient and cost-effective system to prove high yields of virus.

Influenza infection is associated with about 36,000 deaths and more than 200,000 hospitalizations every year in the United States. The continuous ...

Establishment of a High-throughput Setup for Screening Small Molecules That Modulate c-di-GMP Signaling in Pseudomonas aeruginosa

Bacterial resistance to traditional antibiotics has driven research attempts to identify new drug targets in recently discovered regulatory pathways. Regulatory systems that utilize intracellular cyclic di-GMP (c-di-GMP) as a second messenger are one such class of target. c-di-GMP is a signaling molecule found in almost all bacteria that acts to regulate an extensive range of processes including antibiotic resistance, biofilm formation and virulence. The understanding of how c-di-GMP signaling controls aspects of antibiotic resistant biofilm development has suggested approaches whereby alteration of the cellular concentrations of the nucleotide or disruption of these signaling pathways may lead to reduced biofilm formation or increased susceptibility of the biofilms to antibiotics. We describe a simple high-throughput bioreporter protocol, based on green fluorescent protein (GFP), whose expression is under the control of the c-di-GMP responsive promoter cdrA, to rapidly screen for small molecules with the potential to modulate c-di-GMP cellular levels in Pseudomonas aeruginosa (P. aeruginosa). This simple protocol can screen upwards of 3,500 compounds within 48 hours and has the ability to be adapted to multiple microorganisms.

Establishment of a High-throughput Setup for Screening Small Molecules That Modulate c-di-GMP Signaling in Pseudomonas aeruginosa

This article describes the high-throughput assay that has been successfully established to screen large libraries of small molecules for their potential ability to manipulate cellular levels of cyclic di-GMP in Pseudomonas aeruginosa, providing a new powerful tool for antibacterial drug discovery and compound testing.

Bacterial resistance to traditional antibiotics has driven research attempts to identify new drug targets in recently discovered regulatory pathways. ...

Cefoperazone-treated Mouse Model of Clinically-relevant Clostridium difficile Strain R20291

Clostridium difficile is an anaerobic, gram-positive, spore-forming enteric pathogen that is associated with increasing morbidity and mortality and consequently poses an urgent threat to public health. Recurrence of a C. difficile infection (CDI) after successful treatment with antibiotics is high, occurring in 20-30% of patients, thus necessitating the discovery of novel therapeutics against this pathogen. Current animal models of CDI result in high mortality rates and thus do not approximate the chronic, insidious disease manifestations seen in humans with CDI. To evaluate therapeutics against C. difficile, a mouse model approximating human disease utilizing a clinically-relevant strain is needed. This protocol outlines the cefoperazone mouse model of CDI using a clinically-relevant and genetically-tractable strain, R20291. Techniques for clinical disease monitoring, C. difficile bacterial enumeration, toxin cytotoxicity, and histopathological changes throughout CDI in a mouse model are detailed in the protocol. Compared to other mouse models of CDI, this model is not uniformly lethal at the dose administered, allowing for the observation of a prolonged clinical course of infection concordant with the human disease. Therefore, this cefoperazone mouse model of CDI proves a valuable experimental platform to assess the effects of novel therapeutics on the amelioration of clinical disease and on the restoration of colonization resistance against C. difficile.

Cefoperazone-treated Mouse Model of Clinically-relevant Clostridium difficile Strain R20291

This protocol outlines the cefoperazone mouse model of Clostridium difficile infection (CDI) using a clinically-relevant and genetically-tractable strain, R20291. Emphasis on clinical disease monitoring, C. difficile bacterial enumeration, toxin cytotoxicity, and histopathological changes throughout CDI in a mouse model are detailed in the protocol.

Clostridium difficile is an anaerobic, gram-positive, spore-forming enteric pathogen that is associated with increasing morbidity and mortality and ...

An IL-8 Transiently Transgenized Mouse Model for the In Vivo Long-term Monitoring of Inflammatory Responses

Airway inflammation is often associated with bacterial infections and represents a major determinant of lung disease. The in vivo determination of the pro-inflammatory capabilities of various factors is challenging and requires terminal procedures, such as bronchoalveolar lavage and the removal of lungs for in situ analysis, precluding longitudinal visualization in the same mouse. Here, lung inflammation is induced through the intratracheal instillation of Pseudomonas aeruginosa culture supernatant (SN) in transiently transgenized mice expressing the luciferase reporter gene under the control of a heterologous IL-8 bovine promoter. Luciferase expression in the lung is monitored by in vivo bioluminescent image (BLI) analysis over a 2.5- to 48-h timeframe following the instillation. The procedure can be repeated multiple times within 2 - 3 months, thus permitting the evaluation of the inflammatory response in the same mice without the need to terminate the animals. This approach permits the monitoring of pro- and anti-inflammatory factors acting in the lung in real time and appears suitable for functional and pharmacological studies.

An IL-8 Transiently Transgenized Mouse Model for the In Vivo Long-term Monitoring of Inflammatory Responses

The method described here allows for the visualization of IL-8 promoter-dependent inflammation activation in the lungs of mice through non-invasive bioluminescence imaging (BLI). The same animal can be subjected to BLI multiple times for up to two months from the time of delivery of the luciferase reporter construct.

Airway inflammation is often associated with bacterial infections and represents a major determinant of lung disease. The in vivo determination of the ...

Effects of Taste Signaling Protein Abolishment on Gut Inflammation in an Inflammatory Bowel Disease Mouse Model

Inflammatory bowel disease (IBD) is one of the immune-related gastrointestinal disorders, including ulcerative colitis and Crohn's disease, that affects the life quality of millions of people worldwide. IBD symptoms include abdominal pain, diarrhea, and rectal bleeding, which may result from the interactions among gut microbiota, food components, intestinal epithelial cells, and immune cells. It is of particular importance to assess how each key gene expressed in intestinal epithelial and immune cells affects inflammation in the colon. G protein-coupled taste receptors, including G protein subunit &#945;-gustducin and other signaling proteins, have been found in the intestines. Here, we use &#945;-gustducin as a representative and describe a dextran sulfate sodium (DSS)-induced IBD model to evaluate the effect of gustatory gene mutations on gut mucosal immunity and inflammation. This method combines gene knockout technology with the chemically induced IBD model, and thus can be applied to assess the outcome of gustatory gene nullification as well as other genes that may exuberate or dampen the DSS-induced immune response in the colon. Mutant mice are administered with DSS for a certain period during which their body weight, stool, and rectal bleeding are monitored and recorded. At different timepoints during administration, some mice are euthanized, then the sizes and weights of their spleens and colons are measured and gut tissues are collected and processed for histological and gene expression analyses. The data show that the &#945;-gustducin knockout results in excessive weight loss, diarrhea, intestinal bleeding, tissue damage, and inflammation vs. wild-type mice. Since the severity of induced inflammation is affected by mouse strains, housing environment, and diet, optimization of DSS concentration and administration duration in a pilot experiment is particularly important. By adjusting these factors, this method can be applied to assess both anti- and pro-inflammatory effects.

Here we present a protocol to investigate the effect of the nullification of gustation-related genes on immune responses in a dextran sulfate sodium (DSS)-induced inflammatory bowel disease (IBD) mouse model.

Inflammatory bowel disease (IBD) is one of the immune-related gastrointestinal disorders, including ulcerative colitis and Crohn's disease, that affects ...

Multicolor Flow Cytometry-based Quantification of Mitochondria and Lysosomes in T Cells

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular components responsible for supplying cell energy; however, excess mitochondria also produce reactive oxygen species (ROS) that could cause cell death. Therefore, the number of mitochondria must constantly be adjusted to fit the needs of the cells. This dynamic regulation is achieved in part through the function of lysosomes that remove surplus/damaged organelles and macromolecules. Hence, cellular mitochondrial and lysosomal contents are key indicators to evaluate the metabolic adjustment of cells. With the development of probes for organelles, well-characterized lysosome or mitochondria-specific dyes have become available in various formats to label cellular lysosomes and mitochondria. Multicolor flow cytometry is a common tool to profile cell phenotypes, and has the capability to be integrated with other assays. Here, we present a detailed protocol of how to combine organelle-specific dyes with surface markers staining to measure the amount of lysosomes and mitochondria in different T cell populations on a flow cytometer.

This article illustrates a powerful method to quantify mitochondria or lysosomes in living cells. The combination of lysosome- or mitochondria-specific dyes with fluorescently conjugated antibodies against surface markers allows the quantification of these organelles in mixed cell populations, like primary cells harvested from tissue samples, by using multicolor flow cytometry.

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular ...

Propagation of the Microsporidian Parasite Edhazardia aedis in Aedes aegypti Mosquitoes

Edhazardia aedis is a microsporidian parasite of Aedes aegypti mosquitoes, a disease vector that transmits multiple arboviruses which cause millions of disease cases each year. E. aedis causes mortality and reduced reproductive fitness in the mosquito vector and has been explored for its potential as a biocontrol agent. The protocol we present for culturing E. aedis is based on its natural infection cycle, which involves both horizontal and vertical transmission at different life stages of the mosquito host. Ae. aegypti mosquitoes are exposed to spores in the larval stage. These infected larvae then mature into adults and transmit the parasite vertically to their offspring. Infected offspring are then used as a source of spores for future horizontal transmission. Culturing E. aedis can be challenging to the uninitiated given the complexities of the parasite&#8217;s life cycle, and this protocol provides detailed guidance and visual aids for clarification.

Propagation of the Microsporidian Parasite Edhazardia aedis in Aedes aegypti Mosquitoes

A protocol to culture the microsporidian parasite Edhazardia aedis. The parasite is passaged from one generation of Aedes aegypti mosquitoes to the next via horizontal transfer at the larval stage followed by vertical transmission at the adult stage. Live sporoplasms survive long-term in infected eggs.

Edhazardia aedis is a microsporidian parasite of Aedes aegypti mosquitoes, a disease vector that transmits multiple arboviruses which cause millions of ...