<ol style="padding:5px;">
<li>Equipment Set-up<ol style="padding:5px;">
<li>Select an area in the lab where the room temperature is at or above 20&deg;C.</li>
<li>Avoid locations near air conditioning vents and/or loud noise.</li>
<li>Avoid exposing the animal to odors that may cause irritation and/or stress, such as perfume, cologne, chemicals, and cleaners.</li>
<li>Turn on the CODA Controller.</li>
<li>Turn on the Warming Platform and set to Level 3. If room temperature is above 25&deg;C, start on Levels 1 or 2.</li>
<li>Controller Diagnostics Test<ol style="padding:5px;">
<li>Open the CODA Software.</li>
<li>Select the CODA Device by clicking on it.</li>
<li>Click on &ldquo;Test Selected Device&rdquo;.</li>
<li>Select the cuffs and channels to test and click &ldquo;Test&rdquo;.</li>
<li>After the test is complete, close the Device Test window.</li>
</ol></li>
</ol></li>
<li>Software Set-up<ol style="padding:5px;">
<li>Manage Personnel and Animals<ol style="padding:5px;">
<li>Manage Personnel<ol style="padding:5px;">
<li>Select Tools &gt; Manage Personnel.</li>
<li>Click on &ldquo;Researcher 1&rdquo; to change the name.</li>
<li>Click on subsequent rows to add additional researchers.</li>
<li>Click on &ldquo;Technicians Tab&rdquo; to modify or add technicians.</li>
<li>Click on &ldquo;Specimens Tab&rdquo; to name the animal for easy identification and to select the animal type; rat or mouse.</li>
<li>Click on &ldquo;Animals Tab&rdquo; to alter the animal type and sensitivity.</li>
</ol></li>
</ol></li>
<li>New Experiment<ol style="padding:5px;">
<li>Select File &gt; New &gt; Experiment</li>
<li>Begin a New Experiment<ol style="padding:5px;">
<li>Enter the Experiment Name.</li>
<li>Select the Key Researcher.</li>
<li>Select a Begin Date and click &ldquo;Next&rdquo;.</li>
</ol></li>
<li>Basic Session Info<ol style="padding:5px;">
<li>Enter the Session Name</li>
<li>Set &ldquo;Acclimation Cycles&rdquo; to 5. If you prefer not to include acclimation cycles, set to zero. To assist with acclimating the animal to the holder and cuffs, add additional cycles to your experiment and run the experiment for 5 consecutive days.</li>
<li>Set &ldquo;Number of Sets&rdquo; to 1.</li>
<li>Set &ldquo;Time Between Sets&rdquo; to 30 seconds for multiple sets</li>
<li>Set &ldquo;Cycles per Set&rdquo; to 20 cycles.</li>
<li>Set &ldquo;Time Between Cycles&rdquo; to 5 seconds and click &ldquo;Next&rdquo;.</li>
</ol></li>
</ol></li>
<li>Specimen Selection<ol style="padding:5px;">
<li style="list-style-type:none;">&nbsp;<ol style="padding:5px;">
<li>Select the specimen from the specimen pool and assign it to the first available channel and click &ldquo;Next&rdquo;. </li>
</ol></li>
<li>Session Parameters<ol style="padding:5px;">
<li>Set &ldquo;Maximum Occlusion Pressure&rdquo; to 250mmHg.</li>
<li>Set &ldquo;Deflation Time&rdquo; to 20 seconds for mice, 15 seconds for rats.</li>
<li>Set &ldquo;Minimum Volume&rdquo; to 15ul.</li>
<li>Set &ldquo;Display Style&rdquo; to One Channel per Graph.</li>
<li>Add additional Researchers and Technicians and click &ldquo;Next&rdquo;.</li>
</ol></li>
<li>Session Script<ol style="padding:5px;">
<li>Review the Session Script and click &ldquo;Next&rdquo;.</li>
<li>Do not select Finish until you complete the Animal Preparation section.</li>
</ol></li>
</ol></li>
</ol></li>
<li>Animal Preparation<ol style="padding:5px;">
<li>Animal Set-up<ol style="padding:5px;">
<li>Animal Holder<ol style="padding:5px;">
<li>To train and improve animal acclimation, add additional cycles to your experiment and run the experiment for five consecutive days. You also can assist with acclimation by placing the animal in the holder for 15 minutes for 3 consecutive days prior to the actual study.</li>
<li>Place each animal in a holder by picking up the animal by the tail and gently placing the animal into the rear of the holder which faces the open end of the nose cone. Note: the size of animal in relation to the size of the selected holder. The animal should be able to freely enter the holder.</li>
<li>Carefully secure the rear hatch to the holder by turning the red screw cap on the rear hatch. Care should be taken to avoid pinching the tail or any other body parts while securing the rear hatch.</li>
<li>Slide the nose cone toward the rear hatch, limiting the movement of the animal. The nose cone should be in a position to limit the animal from turning around while inside the holder. </li>
<li>Place the holder onto the warming platform in the designated position. If testing the same group of animals over a period of time, be sure to rotate the holder position on the warming platform to improve consistency and accuracy of your BP measurements. Allow the animal at least 5-minutes to acclimate to the holder.</li>
<li>Do not touch or handle the animal while inside the holder.&nbsp; The increased contact could irritate the animal.</li>
<li>Never leave the animal in the holders unattended.</li>
</ol></li>
<li>Cuff Placement<ol style="padding:5px;">
<li>Thread the tail through the &ldquo;Occlusion Cuff.&rdquo; Place this cuff as close to the base of the tail as possible without force.</li>
<li>Secure the &ldquo;Occlusion Cuff Tubing&rdquo; in the notch on the top rear of the holder.</li>
<li>Thread the tail through the &ldquo;VPR Cuff&rdquo;, placing it within 2mm of the &ldquo;Occlusion Cuff&rdquo;.</li>
<li>Secure the &ldquo;VPR Cuff Tubing&rdquo; in the notch on the top rear of the holder.</li>
<li>Attach the cuffs to the CODA Controller.</li>
</ol></li>
<li>Animal Temperature<ol style="padding:5px;">
<li>Allow the animal at least 5-minutes to thermoregulate.</li>
<li>Record the temperature of the animal frequently with an infrared thermometer. The animal temperature should be 32 to 35&deg;C.</li>
</ol></li>
</ol></li>
</ol></li>
<li>Experiment<ol style="padding:5px;">
<li>Begin Experiment<ol style="padding:5px;">
<li>Click &ldquo;Finish&rdquo; and the experiment will begin immediately.</li>
</ol></li>
<li>End Experiment<ol style="padding:5px;">
<li>Animal<ol style="padding:5px;">
<li>Immediately remove the animal from the cuffs and holder.</li>
<li>Return the animal to their cage after the &ldquo;Simple Session Summary Report&rdquo; is displayed. </li>
</ol></li>
<li>Data Collection<ol style="padding:5px;">
<li>After completion of the last cycle the accepted cycles will be automatically displayed in spreadsheet format within the CODA application, click on the &ldquo;Export to Excel&rdquo; button.</li>
<li>Select the folder to place the file in, name the file and click &ldquo;Save&rdquo;.</li>
<li>Open the Excel application and click on &ldquo;Open&rdquo;.</li>
<li>View all file types to display the &ldquo;.csv file saved&rdquo;, click on the file.</li>
</ol></li>
<li>Data Processing<ol style="padding:5px;">
<li>There are several ways to process the data within the Excel file. A common practice is to obtain the average and standard deviation.</li>
<li>Delete measurements if the standard deviation is greater than 30.</li>
<li>Additionally, rejected cycles may be viewed and the entire database can be easily exported to Excel. Consult the User&rsquo;s Manual for additional information.</li>
</ol></li>
</ol></li>
</ol></li>
</ol>

The authors have nothing to disclose.

measuring blood pressure in mice using volume pressure recording, a tail-cuff method

The CODA 8-Channel High Throughput Non-Invasive Blood Pressure system measures the blood pressure in up to 8 mice or rats simultaneously. The CODA tail-cuff system uses Volume Pressure Recording (VPR) to measure the blood pressure by determining the tail blood volume. A specially designed differential pressure transducer and an occlusion tail-cuff measure the total blood volume in the tail without the need to obtain the individual pulse signal. Special attention is afforded to the length of the occlusion cuff in order to derive the most accurate blood pressure readings. VPR can easily obtain readings on dark-skinned rodents, such as C57BL6 mice and is MRI compatible. The CODA system provides you with measurements of six (6) different blood pressure parameters; systolic and diastolic blood pressure, heart rate, mean blood pressure, tail blood flow, and tail blood volume. Measurements can be made on either awake or anesthetized mice or rats. The CODA system includes a controller, laptop computer, software, cuffs, animal holders, infrared warming pads, and an infrared thermometer. There are seven different holder sizes for mice as small as 8 grams to rats as large as 900 grams.

Watch this Scientific Journal Video about Measuring Blood Pressure in Mice using Volume Pressure Recording, a Tail-cuff Method at JoVE.com

Measuring Blood Pressure in Mice using Volume Pressure Recording, a Tail-cuff Method

The CODA 8-Channel High Throughput Non-Invasive Blood Pressure system measures the blood pressure in up to 8 mice or rats simultaneously. This tail-cuff system uses Volume Pressure Recording (VPR) to measure the blood pressure by determining the tail blood volume.

The CODA 8-Channel High Throughput Non-Invasive Blood Pressure system measures the blood pressure in up to 8 mice or rats simultaneously. The CODA ...

measuring-blood-pressure-mice-using-volume-pressure-recording-tail

Research

JoVE Journal

Biology

51.5K 次观看.  University of Kentucky. 尾声，8通道高通量非侵入性血压系统测量血压，同时8只小鼠或大鼠。这尾袖系统采用体积压力记录（VPR）来衡量确定的尾巴血容量血压。

视频: Measuring Blood Pressure in Mice using Volume Pressure Recording, a Tail-cuff Method

Pressure-polishing Pipettes for Improved Patch-clamp Recording

Pressure-polishing is a method for shaping glass pipettes for patch-clamp recording. We first developed this method for fabricating pipettes suitable for recording from small (&lt;3 m) neuronal cell bodies. The basic principal is similar to glass-blowing and combines air pressure and heat to modify the shape of patch pipettes prepared by a conventional micropipette puller. It can be applied to so-called soft (soda lime) and hard (borosilicate) glasses. Generally speaking, pressure polishing can reduce pipette resistance by 25% without decreasing the diameter of the tip opening (Goodman and Lockery, 2000). It can be applied to virtually any type of glass and requires only the addition of a high-pressure valve and fitting to a microforge. This technique is essential for recording from ultrasmall cells (&lt;5 m) and can also improve single-channel recording by minimizing pipette resistance. The blunt shape is also useful for perforated-patch clamp recording since this tip shape results in a larger membrane bleb available for perforation.

This is a guide to modifying the shape of glass micropipettes. Specifically, by using heat and air pressure the taper is widened without increasing the tip opening, leading to lower pipette resistance. This is critical to obtain low noise recordings of small cells but is useful in many applications.

Pressure-polishing is a method for shaping glass pipettes for patch-clamp recording.  We first developed this method for fabricating pipettes suitable for ...

Making Gynogenetic Diploid Zebrafish by Early Pressure

Heterozygosity in diploid eukaryotes often makes genetic studies cumbersome. Methods that produce viable homozygous diploid offspring directly from heterozygous females allow F1 mutagenized females to be screened directly for deleterious mutations in an accelerated forward genetic screen. Streisinger et al.1,2 described methods for making gynogenetic (homozygous) diploid zebrafish by activating zebrafish eggs with ultraviolet light-inactivated sperm and preventing either the second meiotic or the first zygotic cell division using physical treatments (heat or pressure) that deploymerize microtubules. The "early pressure" (EP) method blocks the meiosis II, which occurs shortly after fertilization. The EP method produces a high percentage of viable embryos that can develop to fertile adults of either sex. The method generates embryos that are homozygous at all loci except those that were separated from their centromere by recombination during meiosis I. Homozygous mutations are detected in EP clutches at between 50% for centromeric loci and less than 1% for telomeric loci. This method is reproduced verbatim from the Zebrafish Book3.

This is a method for generating gynogenetic diploid zebrafish embryos (embryos whose only genetic contribution comes from the mother) by blocking the second meiotic division immediately after fertilization with ultraviolet light-inactivated sperm. EP embryos are not fully homozygous due to recombination during the first meiotic division, however they are homozygous at all loci that have not been separated from their centromere by recombination.

Heterozygosity in diploid eukaryotes often makes genetic studies cumbersome. Methods that produce viable homozygous diploid offspring directly from ...

Ambulatory ECG Recording in Mice

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although the surface ECG is utilized for short-term measurements of waveform intervals, it is not practical for long-term studies of heart rate variability or the capture of rare episodes of arrhythmias. Implantable ECG telemeters offer the advantages of simple surgical implantation, long-term recording of electrograms in ambulatory mice, and scalability with simultaneous recordings of multiple animals. Here, we present a step-by-step guide to the implantation of telemeters for ambulatory ECG recording in mice. Careful adherence to aseptic technique is required for favorable survival results with the possibility of implantation and recording from weeks to months. Thus, implantable ECG telemetry is a valuable tool for detection of critical information on cardiac electrophysiology in ambulatory animal models such as the mouse. 

Telemetric ECG has emerged as an essential tool in evaluating animal models for cardiac arrhythmias and sudden cardiac death. Here, we present a stepwise guide to telemetric ECG recordings for application in long-term ambulatory ECG monitoring in mice.

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although ...

Isolation and Culture of Pulmonary Endothelial Cells from Neonatal Mice

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells (HUVECs) have shown to be convenient, easy to obtain and culture, and thus are the most widely studied endothelial cells. However, for research focused on processes like angiogenesis, permeability or many others, microvascular endothelial cells (ECs) are a much more physiologically relevant model to study 2. Furthermore, ECs isolated from knockout mice provide a useful tool for analysis of protein function ex vivo. Several approaches to isolate and culture microvascular ECs of different origin have been reported to date 3-7, but consistent isolation and culture of pure ECs is still a major technical problem in many laboratories. Here, we provide a step-by-step protocol on a reliable and relatively simple method of isolating and culturing mouse lung endothelial cells (MLECs). In this approach, lung tissue obtained from 6- to 8-day old pups is first cut into pieces, digested with collagenase/dispase (C/D) solution and dispersed mechanically into single-cell suspension. MLECS are purified from cell suspension using positive selection with anti-PECAM-1 antibody conjugated to Dynabeads using a Magnetic Particle Concentrator (MPC). Such purified cells are cultured on gelatin-coated tissue culture (TC) dishes until they become confluent. At that point, cells are further purified using Dynabeads coupled to anti-ICAM-2 antibody. MLECs obtained with this protocol exhibit a cobblestone phenotype, as visualized by phase-contrast light microscopy, and their endothelial phenotype has been confirmed using FACS analysis with anti-VE-cadherin 8 and anti-VEGFR2 9 antibodies and immunofluorescent staining of VE-cadherin. In our hands, this two-step isolation procedure consistently and reliably yields a pure population of MLECs, which can be further cultured. This method will enable researchers to take advantage of the growing number of knockout and transgenic mice to directly correlate in vivo studies with results of in vitro experiments performed on isolated MLECs and thus help to reveal molecular mechanisms of vascular phenotypes observed in vivo.

Here, we describe a protocol for isolation and culture of murine pulmonary endothelial cells. This method comprises mechanic and enzymatic lung tissue dissociation as well as a 2-step purification process using anti-PECAM-1 and anti-ICAM-2 antibodies conjugated to magnetic beads, which produces a pure endothelial cell population of mostly microvascular origin.

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells ...

Assessing Murine Resistance Artery Function Using Pressure Myography

Pressure myograph systems are exquisitely useful in the functional assessment of small arteries, pressurized to a suitable transmural pressure. The near physiological condition achieved in pressure myography permits in-depth characterization of intrinsic responses to pharmacological and physiological stimuli, which can be extrapolated to the in vivo behavior of the vascular bed. Pressure myograph has several advantages over conventional wire myographs. For example, smaller resistance vessels can be studied at tightly controlled and physiologically relevant intraluminal pressures. Here, we study the ability of 3rd order mesenteric arteries (3-4 mm long), preconstricted with phenylephrine, to vaso-relax in response to acetylcholine. Mesenteric arteries are mounted on two cannulas connected to a pressurized and sealed system that is maintained at constant pressure of 60 mmHg. The lumen and outer diameter of the vessel are continuously recorded using a video camera, allowing real time quantification of the vasoconstriction and vasorelaxation in response to phenylephrine and acetylcholine, respectively.
	To demonstrate the applicability of pressure myography to study the etiology of cardiovascular disease, we assessed endothelium-dependent vascular function in a murine model of systemic hypertension. Mice deficient in the &alpha;1 subunit of soluble guanylate cyclase (sGC&alpha;1-/-) are hypertensive when on a 129S6 (S6) background (sGC&alpha;1-/-S6) but not when on a C57BL/6 (B6) background (sGC&alpha;1-/-B6). Using pressure myography, we demonstrate that sGC&alpha;1-deficiency results in impaired endothelium-dependent vasorelaxation. The vascular dysfunction is more pronounced in sGC&alpha;1-/-S6 than in sGC&alpha;1-/-B6 mice, likely contributing to the higher blood pressure in sGC&alpha;1-/-S6 than in sGC&alpha;1-/-B6 mice. 
	Pressure myography is a relatively simple, but sensitive and mechanistically useful technique that can be used to assess the effect of various stimuli on vascular contraction and relaxation, thereby augmenting our insight into the mechanisms underlying cardiovascular disease.

In pressure myography, an intact small segment of a vessel is mounted onto two small cannulas and pressurized to a suitable luminal pressure. Here, we describe the method to measure vasorelaxation response of the mouse 3rd order mesenteric arteries in c57 and sGC&alpha;1-/- mice using pressure myography.

Pressure myograph systems are  exquisitely useful in the functional assessment of small arteries, pressurized  to a suitable transmural pressure. The  ...

One-channel Cell-attached Patch-clamp Recording

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate depends on properties intrinsic to each channel protein as well as the mechanism by which it is generated and controlled and represents an important area of current research. Information about the operational dynamics of ion channel proteins can be obtained by observing long stretches of current produced by a single molecule. Described here is a protocol for obtaining one-channel cell-attached patch-clamp current recordings for a ligand gated ion channel, the NMDA receptor, expressed heterologously in HEK293 cells or natively in cortical neurons. Also provided are instructions on how to adapt the method to other ion channels of interest by presenting the example of the mechano-sensitive channel PIEZO1. This method can provide data regarding the channel&#8217;s conductance properties and the temporal sequence of open-closed conformations that make up the channel&#8217;s activation mechanism, thus helping to understand their functions in health and disease.

Described here is a procedure for obtaining long stretches of current recording from one ion channel with the cell-attached patch-clamp technique. This method allows for observing, in real time, the pattern of open-close channel conformations that underlie the biological signal. These data inform about channel properties in undisturbed biological membranes.

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate ...

Measuring Respiratory Function in Mice Using Unrestrained Whole-body Plethysmography

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative assessment of lung function in rodent models is an important tool in the development of future therapies. Commonly used techniques for assessing respiratory function including invasive plethysmography and forced oscillation. While these techniques provide valuable information, data collection can be fraught with artefacts and experimental variability due to the need for anesthesia and/or invasive instrumentation of the animal. In contrast, unrestrained whole-body plethysmography (UWBP) offers a precise, non-invasive, quantitative way by which to analyze respiratory parameters. This technique avoids the use of anesthesia and restraints, which is common to traditional plethysmography techniques. This video will demonstrate the UWBP procedure including the equipment set up, calibration and lung function recording. It will explain how to analyze the collected data, as well as identify experimental outliers and artefacts that results from animal movement. The respiratory parameters obtained using this technique include tidal volume, minute volume, inspiratory duty cycle, inspiratory flow rate and the ratio of inspiration time to expiration time. UWBP does not rely on specialized skills and is inexpensive to perform. A key feature of UWBP, and most appealing to potential users, is the ability to perform repeated measures of lung function on the same animal.

The assessment of respiratory physiology has traditionally relied upon techniques, which require restraint or sedation of the animal. Unrestrained whole-body plethysmography, however, provides precise, non-invasive, quantitative analysis of respiratory physiology in animal models. In addition, the technique allows repeated respiratory assessment of mice allowing for longitudinal studies.

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative ...

A Method for Screening and Validation of Resistant Mutations Against Kinase Inhibitors

The discovery of BCR/ABL as a driver oncogene in chronic myeloid leukemia (CML) resulted in the development of Imatinib, which, in fact, demonstrated the potential of targeting the kinase in cancers by effectively treating the CML patients. This observation revolutionized drug development to target the oncogenic kinases implicated in various other malignancies, such as, EGFR, B-RAF, KIT and PDGFRs. However, one major drawback of anti-kinase therapies is the emergence of drug resistance mutations rendering the target to have reduced or lost affinity for the drug. Understanding the mechanisms employed by resistant variants not only helps in developing the next generation inhibitors but also gives impetus to clinical management using personalized medicine. We reported a retroviral vector based screening strategy to identify the spectrum of resistance conferring mutations in BCR/ABL, which has helped in developing the next generation BCR/ABL inhibitors. Using Ruxolitinib and JAK2 as a drug target pair, here we describe in vitro screening methods that utilizes the mouse BAF3 cells expressing the random mutation library of JAK2 kinase.

Emergence of genetic resistance against kinase inhibitor therapy poses significant challenge for effective cancer therapy. Identification and characterization of resistant mutations against a newly developed drug helps in better clinical management and next generation drug design. Here, we describe our protocol for in vitro screening and validation of resistant mutations.

The discovery of BCR/ABL as a driver oncogene in chronic myeloid leukemia (CML) resulted in the development of Imatinib, which, in fact, demonstrated the ...

Cardiac Pressure-Volume Loop Analysis Using Conductance Catheters in Mice

Cardiac pressure-volume loop analysis is the &#8220;gold-standard&#8221; in the assessment of load-dependent and load-independent measures of ventricular systolic and diastolic function. Measures of ventricular contractility and compliance are obtained through examination of cardiac response to changes in afterload and preload. These techniques were originally developed nearly three decades ago to measure cardiac function in large mammals and humans. The application of these analyses to small mammals, such as mice, has been accomplished through the optimization of microsurgical techniques and creation of conductance catheters. Conductance catheters allow for estimation of the blood pool by exploiting the relationship between electrical conductance and volume. When properly performed, these techniques allow for testing of cardiac function in genetic mutant mouse models or in drug treatment studies. The accuracy and precision of these studies are dependent on careful attention to the calibration of instruments, systematic conduct of hemodynamic measurements and data analyses. We will review the methods of conducting pressure-volume loop experiments using a conductance catheter in mice.

Cardiac pressure-volume loop analysis is the most comprehensive way to measure cardiac function in the intact heart. We describe a technique to perform and analyze cardiac pressure volume loops, using conductance catheters.

Cardiac pressure-volume loop analysis is the &#8220;gold-standard&#8221; in the assessment of load-dependent and load-independent measures of ventricular ...

Measuring Pressure Volume Loops in the Mouse

Understanding the causes and progression of heart disease presents a significant challenge to the biomedical community. The genetic flexibility of the mouse provides great potential to explore cardiac function at the molecular level. The mouse&#39;s small size does present some challenges in regards to performing detailed cardiac phenotyping. Miniaturization and other advancements in technology have made many methods of cardiac assessment possible in the mouse. Of these, the simultaneous collection of pressure and volume data provides a detailed picture of cardiac function that is not available through any other modality. Here a detailed procedure for the collection of pressure-volume loop data is described. Included is a discussion of the principles underlying the measurements and the potential sources of error. Anesthetic management and surgical approaches are discussed in great detail as they are both critical to obtaining high quality hemodynamic measurements. The principles of hemodynamic protocol development and relevant aspects of data analysis are also addressed.

This manuscript describes a detailed protocol for the collection of pressure-volume data from the mouse.

Understanding the causes and progression of heart disease presents a significant challenge to the biomedical community. The genetic flexibility of the ...