This research was funded by Guangdong Provincial Key Laboratory of Drug Non-clinical Evaluation and Research (No.2018B030323024) and Key Program &#34;New Drug Creation&#34; of Guangdong Key Research and Development Plan (No.2019B020202001), Guangzhou Fundamental and Application Foundation Research Project (No.202002030249 and No.202002030156).

All animal experiments described were approved by the Institutional Animal Care and Use Committee (IACUC) of Guangdong Lewwin Pharmaceutical Research Institute Co., Ltd. Male and female Sprague Dawley (SD) rats, ~6-11 weeks were used for the experiments. The rats were reared following the guidelines for the care and use of laboratory animals13.
1. Animal preparation
NOTE: All the SD rats used in this study were awake and were not anesthetized/euthanized. The skill of restraint handling by grasping the skin on the back of the animal is needed.
<ol>
	<li>Find the lowest point of the subclavian triangular fossa between the neck and the forelimb in a rat. Move ~2-3 mm toward the head and reach the needlepoint at the blood collection site (Figure 1). Remove the hair using an electric shaver. 
	NOTE: Depending on the experimental needs, serum or plasma may be required. Plasma is taken as an example in this protocol, and it will not affect the operation of blood collection.</li>
	<li>Prepare cotton swabs stained with 75% alcohol for wiping and disinfecting and dry cotton swabs for wiping.</li>
</ol>
2. Sampling of blood
NOTE: At least 2 people, both of whom must be experienced in blood collection and rat restraint techniques should carry out these steps.
<ol>
	<li>Grasp the skin on the back of the neck with one hand to keep the rat&#39;s head, neck, and breast upright and expose the injection site (Video 1). Straighten the forelimb on the side of the injection site to maintain its level.</li>
	<li>Keep the syringe parallel to the rat&#39;s head with the other hand and tilt the syringe outward 5&#176;-10&#176; so that the tip is tilted toward the ventral direction.</li>
	<li>Insert the needle fully into the anterior cavity. Draw back the syringe to maintain negative pressure within the tube.</li>
	<li>Slowly move the needle from deep to shallow and back up the same path. When blood has entered the syringe needle, fix the position of the needle (Video 2). 
	NOTE: The blood will then be filled at a constant rate in the syringe, as shown in Figure 2A (front view) and Figure 2B (side view).</li>
	<li>Monitor the maximum blood quantity in accordance with the standards set by the institution&#8217;s animal care and use committee. This depends on the weight and health of the animal. In absence of other requirements, do&#160;not remove more than 20% of the animal&#39;s total blood volume within 24 h, which requires ~3 weeks of recuperation14.</li>
	<li>When sufficient blood sample is collected, immediately withdraw the syringe and prepare for blood treatment (step 3).</li>
	<li>Pressurize the rat&#39;s injection site for ~1-2 min to stop the bleeding. Since the collection site is in the lower part of the neck, pinch the skin of the subclavian vein to stop the bleeding by pressing on it (Figure 3). 
	NOTE: Blood can be collected alternately from the bilateral subclavian vein when repeated blood collection is needed.</li>
	<li>Return the rat to the cage and observe its condition.</li>
</ol>
3. Processing the blood sample
<ol>
	<li>Remove the needle from the syringe and discard it in the sharp tool container. Slowly transfer the blood from the syringe to a 1.5 mL tube. Press the syringe against the wall to avoid bubble formations, if any. 
	NOTE: As pressure may cause red blood cells to rupture, remove the needle to prevent hemolysis14.</li>
	<li>Cover the microcentrifuge tube, gently flick it, and turn it upside down at least five times to thoroughly mix the blood with the anticoagulant.</li>
	<li>Centrifuge the whole blood sample at 2000 x g for ~10-15 min at room temperature within 120 min after collecting the plasma.</li>
	<li>Use a pipette gun (see Table of Materials) to transfer the upper plasma into a blank microcentrifuge tube. Do not touch the pipette head with the lower whole blood. Dispose of or re-centrifuge the plasma contaminated with the red blood cells.</li>
	<li>Use the samples immediately or store them at -30 &#176;C. 
	NOTE: The characteristics of the drug determine the storage time. The plasma specimens obtained from the subclavian vein are translucent and light yellow. Hemolysis may turn the plasma red.</li>
</ol>

<ol>
	<li>Shravya, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=International+conference+on+harmonization+of+technical+requirements+for+registration+of+pharmaceuticals+for+human+use.+ICH+M2+EWG.">International conference on harmonization of technical requirements for registration of pharmaceuticals for human use. ICH M2 EWG.</a> Electronic common technical document. , (2014).</li><li>Tan, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The China Food and Drug Administration (CFDA). , (2015).</li><li>McGuill, M. W., Andrew, N. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biological+effects+of+blood+loss:+Implications+for+sampling+volumes+and+techniques.">Biological effects of blood loss: Implications for sampling volumes and techniques.</a> ILAR Journal. 31 (4), 5-20 (1989).</li><li>Aguilar-Mariscal, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oral+pharmacokinetics+of+meloxicam+in+the+rat+using+a+high-performance+liquid+chromatography+method+in+micro-whole-blood+samples.">Oral pharmacokinetics of meloxicam in the rat using a high-performance liquid chromatography method in micro-whole-blood samples.</a> Methods and Findings in Experimental and Clinical Pharmacology. 29 (9), 587-592 (2007).</li><li>Korfmacher, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Utility+of+capillary+microsampling+for+rat+pharmacokinetic+studies:+Comparison+of+tail-vein+bleed+to+jugular+vein+cannula+sampling.">Utility of capillary microsampling for rat pharmacokinetic studies: Comparison of tail-vein bleed to jugular vein cannula sampling.</a> Journal of Pharmacological and Toxicological Methods. 76, 7-14 (2015).</li><li>JoVE. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=JoVE+Science+Education+Database.+Blood+Withdrawal+I.">JoVE Science Education Database. Blood Withdrawal I.</a> JoVE. , (2021).</li><li>Yang, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subclavian+vein+puncture+as+an+alternative+method+of+blood+sample+collection+in+rats.">Subclavian vein puncture as an alternative method of blood sample collection in rats.</a> Journal of Visualized Experiments. (141), e58499 (2018).</li><li>Zou, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Repeated+blood+collection+from+tail+vein+of+non-anesthetized+rats+with+a+vacuum+blood+collection+system.">Repeated blood collection from tail vein of non-anesthetized rats with a vacuum blood collection system.</a> Journal of Visualized Experiments. (130), e55852 (2017).</li><li>Parasuraman, S., Raveendran, R., Kesavan, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blood+sample+collection+in+small+laboratory+animals.">Blood sample collection in small laboratory animals.</a> Journal of Pharmacology &amp; Pharmacotherapeutics. 1 (2), 87-93 (2010).</li><li>Itziar, F., Arantza, P., Nahia, D., Virginia, P., Juan, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clinical+biochemistry+parameters+in+C57BL/6J+mice+after+blood+collection+from+the+submandibular+vein+and+retroorbital+plexus.">Clinical biochemistry parameters in C57BL/6J mice after blood collection from the submandibular vein and retroorbital plexus.</a> Journal of the American Association for Laboratory Animal Science. 49 (2), 202-206 (2010).</li><li>Heimann, M., Roth, D. R., Ledieu, D., Pfister, R., Classen, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sublingual+and+submandibular+blood+collection+in+mice:+A+comparison+of+effects+on+body+weight,+food+consumption+and+tissue+damage.">Sublingual and submandibular blood collection in mice: A comparison of effects on body weight, food consumption and tissue damage.</a> Laboratory Animals. 44 (4), 352-358 (2010).</li><li>Wren-Dail, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+isoflurane+anesthesia+on+circadian+metabolism+and+physiology+in+rats.">Effect of isoflurane anesthesia on circadian metabolism and physiology in rats.</a> Comparative Medicine. 67 (2), 138-146 (2017).</li><li>National Institute of Health. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Guide for the Care and Use of Laboratory Animals. 8th edition. , (2011).</li><li>Lee, G., Goosens, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sampling+blood+from+the+lateral+tail+vein+of+the+rat.">Sampling blood from the lateral tail vein of the rat.</a> Journal of Visualized Experiments. (99), e52766 (2015).</li><li>Diehl, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+good+practice+guide+to+the+administration+of+substances+and+removal+of+blood,+including+routes+and+volumes.">A good practice guide to the administration of substances and removal of blood, including routes and volumes.</a> Journal of Applied Toxicology. 21 (1), 15-23 (2001).</li><li>Casal, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+and+physiological+methods+of+evaluating+median+nerve+regeneration+in+the+rat.">Functional and physiological methods of evaluating median nerve regeneration in the rat.</a> Journal of Visualized Experiments. (158), e59767 (2020).</li></ol>

Limei Wang, Jianmin Guo, Xiaoman Zhong, Yali Sheng, Qiwen Lai, Hui Song, and Wei Yang have a financial interest in Guangdong Lewwin Pharmaceutical Research Institute Co., Ltd, which, however, did not support this work. The other authors declare no competing interests.

There are certain benefits of collecting blood from the subclavian veins. (1) As the site of the blood collection is readily dissociated, and the venous plexus is not regular due to the different postures of the rats, the described method can easily localize the position of the venous plexus while maintaining the stable and comfortable postures of the rats. (2) The operation is easy and favorable to the rapid development of technicians' skills and less pain to animals. (3) An operating mode to make the animal comfortable...

According to the International Conference on Harmonization of Technical Requirements for Registration of Pharmaceuticals for Human Use (ICH) guidelines1 and the National Medical Products Administration (NMPA) guidelines2, the number of blood collection time points of rats in the toxicokinetic (TK) study needs to meet the requirements of the dynamic drug exposure assessment. The approximate total blood volume of a rat is 55-70 mL/kg of body weight3. The collection time points are generally intensive within 30 min after dosing and decrease after that, and more than ten blood samples need to be collected within 48 h in routine testing4. For example, blood samples are collected at 12-time points (0 min, 5 min, 10 min, 15 min, 30 min, 45 min, 1 h, 2 h, 3 h, 4 h, 8 h, and 12 h) in TK study of orally administered drugs. Researchers must repeatedly collect 200-250 &#181;L of blood in rats to obtain high-quality plasma for the TK testing5.
The blood collection sites in rats include tail blood vessels, retro-orbital plexus vein, submandibular vein, heart, abdominal aorta6,&#160;and so on. Among them, blood collection from the caudal vein of rats is a frequently used method, which requires experienced and skilled operators7,8. To collect blood from the retro-orbital plexus vein is less complicated; however, this method is not recommended as it may damage the rats&#39; eyesight9, and blood from the heart and abdominal aorta is only appropriate for final blood sampling10. Another method of collecting blood from the submandibular vein in a conscious rat has been shown to result in more complications and revealed insufficient blood sample quality11. Therefore, researchers may anesthetize the animal to reduce the difficulty of sampling. Still, the anesthesia also increases the cost of the experiment, and more seriously, it will affect the metabolic state of rats12. The present protocol uses a quick and straightforward method of collecting blood in the subclavian veins of rats without anesthesia, allowing accurate positioning and bilateral alternating blood collection to obtain high-quality samples in a timely and repeated manner.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1.0 mL syringe (with needle, 26 G, 0.45 mm x 15.5 mm)</td>
			<td>Jiangxi Hongda Medical Equipment Group LTD.(Nanchang, Jiangxi Province, China)</td>
			<td>20210629</td>
			<td></td>
		</tr>
		<tr>
			<td>75% alcohol</td>
			<td>Shandong Lierkang Medical Technology Co., Ltd. (Dezhou, Shandong Province, China)</td>
			<td>210717</td>
			<td></td>
		</tr>
		<tr>
			<td>Animal source</td>
			<td>Hunan SJA Laboratory Animal Co., Ltd.</td>
			<td>grade: SPF</td>
			<td>laboratory animal production license number: SCXK (Hunan) 2019-0004, laboratory animal quality certificate number: 4307272101972847</td>
		</tr>
		<tr>
			<td>Cotton swab</td>
			<td>Caoxian Hualu Sanitary Materials Co., Ltd.(Heze, Shandong Province, China)</td>
			<td>20210301</td>
			<td>Need to be sterilized.</td>
		</tr>
		<tr>
			<td>Electric shaver</td>
			<td>Shenzhen Codos Electric Appliance Co. , Ltd.(Shenzhen, Guangdong Province, China)</td>
			<td>CP-6800</td>
			<td></td>
		</tr>
		<tr>
			<td>EP tube, 1.5 mL</td>
			<td>Genetimes ExCell Technology,Inc.(Shanghai, China)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin sodium (2 mL:12500 IU)</td>
			<td>Tianjin biochem pharmaceutical Co.,Ltd(Tianjing, China)</td>
			<td>51200702</td>
			<td>Prepare 1250 IU/mL heparin sodium solution. Add heparin sodium solution into the EP tube in advance and make it evenly distributed on the wall of the EP tube, and dry it at 60&#176;C for use.&#160;</td>
		</tr>
		<tr>
			<td>Labtip (volume range 5-200 &#956;L)</td>
			<td>Thermo Fisher Scientific Oy</td>
			<td>94300120</td>
			<td></td>
		</tr>
		<tr>
			<td>Low speed refrigerated centrifuge</td>
			<td>Hunan Xiangyi Laboratory Instrument Development Co., Ltd.(Changsha, Hunan Province, China)</td>
			<td>L535R</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette gun (20-200 &#956;L)</td>
			<td>BRAND</td>
			<td>12N92305</td>
			<td></td>
		</tr>
		<tr>
			<td>Rats (SD)</td>
			<td>Hunan SJA Laboratory Animal Co., Ltd. (Changsha, Hunan Province, China)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sharp tool container</td>
			<td>Taizhou Huangyan Yikang Plastic Factory (Taizhou, Zhejiang Province, China)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Eye drop</td>
			<td></td>
			<td></td>
			<td>This reagent is not a commodity and the manufacturer requires it to be tested. In the principle of confidentiality, the manufacturer and model cannot be provided.</td>
		</tr>
	</tbody>
</table>

repetitive blood sampling from the subclavian vein of conscious rat

Rats are widely used in pharmacokinetics (PK) and toxicokinetic (TK) studies that need to collect a certain amount of blood at specific time points to detect drug exposure. The rat blood sampling method determines the quality of the plasma and further affects the precision of the test results. The subclavian vein blood collection method described in this protocol collects blood samples repeatedly in the conscious state of animals to meet the needs of PK and TK tests. The skills of restraint handling and appropriate procedure of needle incision ensure the success rate of blood collection. It is easy to operate while ensuring the quality of plasma and at the same time catering to animal welfare. However, this method requires skilled operation, and an improper one may cause animal weakness, pain, lameness, and even mortality. The current method has been used in the testing facility for a 4-week oral toxicity study in Sprague Dawley (SD) rats with TK. The maximum amount of blood collected within 24 h did not exceed 20% of the animal's total blood. The animals' body weight was more than 200 g for males and females. The data showed that the animals' bodyweight increased steadily every week, and the clinical observation was normal after repetitive sample collection.

Good plasma samples from the subclavian vein were translucent pale yellow (Figure 4, the left tube). Improper blood collection or manipulation resulted in hemolysis (Figure 4, the right tube).
The data of the testing facility showed that in a 4-week oral toxicity study of an eye drop in SD rats with TK, blood samples were collected twice at 9-time points (0 h, 0.167 h, 0.5 h, 1 h, 2 h, 4 h, 8 h, 12 h, and 24 h) between the first (Day ...

Watch this Scientific Journal Video about Repetitive Blood Sampling from the Subclavian Vein of Conscious Rat at JoVE.com

Repetitive Blood Sampling from the Subclavian Vein of Conscious Rat

The present protocol describes a simple and efficient method for collecting blood from the subclavian vein in rats. It enables rapid, timely, and easily identifiable sampling without anesthesia and obtains high-quality blood through repetitive sample collection.

Rats are widely used in pharmacokinetics (PK) and toxicokinetic (TK) studies that need to collect a certain amount of blood at specific time points to ...

repetitive-blood-sampling-from-the-subclavian-vein-of-conscious-rat

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JoVE Journal

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4.2K Views.  Guangdong Engineering Research Center for Innovative Drug Evaluation and Research. The present protocol describes a simple and efficient method for collecting blood from the subclavian vein in rats. It enables rapid, timely, and easily identifiable sampling without anesthesia and obtains high-quality blood through repetitive sample collection.

Video: Repetitive Blood Sampling from the Subclavian Vein of Conscious Rat

Hyperinsulinemic-Euglycemic Clamp in the Conscious Rat

Type 2 diabetes (T2D) is rapidly rising in prevalence. Characterized by either inadequate insulin production or the inability to utilize insulin produced, T2D results in elevated blood glucose levels. The "gold-standard" in assessing insulin sensitivity is a hyperinsulinemic-euglycemic clamp or insulin clamp. In this procedure, insulin is infused at a constant rate resulting in a drop in blood glucose. To maintain blood glucose at a constant level, exogenous glucose (D50) is infused into the venous circulation. The amount of glucose infused to maintain homeostasis is indicative of insulin sensitivity. Here, we show the basic clamp procedure in the chronically catheterized, unrestrained, conscious rat. This model allows blood to be collected with minimal stress to the animal. Following the induction of anesthesia, a midline incision is made and the left common carotid artery and right jugular vein are catheterized. Inserted catheters are flushed with heparinized saline, then exteriorized and secured. Animals are allowed to recover for 4-5 days prior to experiments, with weight gain monitored daily. Only those animals who regain weight to pre-surgery levels are used for experiments. On the day of the experiment, rats are fasted and connected to pumps containing insulin and D50. Baseline glucose is assessed from the arterial line and used a benchmark throughout the experiment (euglycemia). Following this, insulin is infused at a constant rate into the venous circulation. To match the drop in blood glucose, D50 is infused. If the rate of D50 infusion is greater than the rate of uptake, a rise in glucose will occur. Similarly, if the rate is insufficient to match whole body glucose uptake, a drop will occur. Titration of glucose continues until stable glucose readings are achieved. Glucose levels and glucose infusion rates during this stable period are recorded and reported. Results provide an index of whole body insulin sensitivity. The technique can be refined to meet specific experimental requirements. It is further enhanced by the use of radioactive tracers that can determine tissue specific insulin-stimulated glucose uptake as well as whole body glucose turnover.

The hyperinsulinemic-euglycemic clamp is the "gold standard" for the assessment of insulin action. Insulin is infused at a constant rate stimulating glucose uptake. The amount of exogenous glucose infused to counter this drop is indicative of insulin sensitivity. Here the procedure is performed on a conscious, unrestrained rat.

Type 2 diabetes (T2D) is rapidly rising in prevalence. Characterized by either inadequate insulin production or the inability to utilize insulin produced, ...

Femoral Arterial and Venous Catheterization for Blood Sampling, Drug Administration and Conscious Blood Pressure and Heart Rate Measurements

In multiple fields of study, access to the circulatory system in laboratory studies is necessary. Pharmacological studies in rats using chronically implanted catheters permit a researcher to effectively and humanely administer substances, perform repeated blood sampling and assists in conscious direct measurements of blood pressure and heart rate. Once the catheter is implanted long-term sampling is possible. Patency and catheter life depends on multiple factors including the lock solution used, flushing regimen and catheter material. This video will demonstrate the methodology of femoral artery and venous catheterization of the rat. In addition the video will demonstrate the use of the femoral venous and arterial catheters for blood sampling, drug administration and use of the arterial catheter in taking measurements of blood pressure and heart rate in a conscious freely-moving rat. A tether and harness attached to a swivel system will allow the animal to be housed and have samples taken by the researcher with minimal disruption to the animal. To maintain patency of the catheter, careful daily maintenance of the catheter is required using lock solution (100 U/ml heparinized saline), machine-ground blunt tip syringe needles and the use of syringe filters to minimize potential contamination. With careful aseptic surgical techniques, proper catheter materials and careful catheter maintenance techniques, it is possible to sustain patent catheters and healthy animals for long periods of time (several weeks).

Chronic catheterization of blood vessels in the rat is often required for administration of substances, obtain blood sample over a period of time or for direct conscious blood pressure measurements. Femoral arterial catheterization of the rat and corresponding measurements of blood pressure in the conscious animal will be demonstrated.

In multiple fields of study, access to the circulatory system in laboratory studies is necessary. Pharmacological studies in rats using chronically ...

Catheterization of the Carotid Artery and Jugular Vein to Perform Hemodynamic Measures, Infusions and Blood Sampling in a Conscious Rat Model

The success of a small animal model to study critical illness is, in part, dependent on the ability of the model to simulate the human condition. Intra-tracheal inoculation of a known amount of bacteria has been successfully used to reproduce the pathogenesis of pneumonia which then develops into sepsis. Monitoring hemodynamic parameters and providing standard clinical treatment including infusion of antibiotics, fluids and drugs to maintain blood pressure is critical to simulate routine supportive care in this model but to do so requires both arterial and venous vascular access. The video details the surgical technique for implanting carotid artery and common jugular vein catheters in an anesthetized rat. Following a 72 hr recovery period, the animals will be re-anesthetized and connected to a tether and swivel setup attached to the rodent housing which connects the implanted catheters to the hemodynamic monitoring system. This setup allows free movement of the rat during the study while continuously monitoring pressures, infusing fluids and drugs (antibiotics, vasopressors) and performing blood sampling.

Vascular accesses to measure hemodynamics, provide fluids and perform blood sampling are important to any small animal model study. We present a technique for implanting catheters into the carotid artery and the common jugular vein in an anesthetized rat for connecting to a system to perform monitoring, infusions and sampling.

The success of a small animal model to study critical illness is, in part, dependent on the ability of the model to simulate the human condition. ...

Repeated Blood Collection from Tail Vein of Non-Anesthetized Rats with a Vacuum Blood Collection System

Blood can be collected from rats in a number of sampling locations. For instance, the tail vein is a superior location for blood sampling. However, the tail vein is thin so that it is sometimes hard to puncture. In addition, the tail vein has low blood flow and requires a long sampling time to get sufficient blood. The present report describes a simple blood sampling method, the vacuum blood collection method, which is usually used to obtain blood samples from patients, here used for non-anesthetized rats. The 22 G butterfly needle tip was inserted into one of the lateral tail veins approximately 2-3 cm from the tip of the tail at an angle of approximately 20&#176;, and blood was collected in the vacuum collection tube by inserting the rubber end of the butterfly needle into the vacuum blood collection tube. The present experimental results show that the success rate was 95% in the experimental group and 90% in the beginner group. The success rate and puncture times were similar between two groups. The sampling duration was significantly shorter in the experimental group compared to beginner group. In conclusion, this vacuum blood collection method for sequential blood sampling from the tail vein of non- anesthetized rats is feasible and easy-to-learn, which might serve as a reliable alternative to other conventionally used blood sampling techniques for rats.

Here, we describe a simple tail vein blood sampling method in non-anesthetized rats using a vacuum extraction tube system. This method reduces the risk of direct exposure to blood and simplifies taking multiple samples from a single venipuncture.

Blood can be collected from rats in a number of sampling locations. For instance, the tail vein is a superior location for blood sampling. However, the ...

Sampling Strategies and Processing of Biobank Tissue Samples from Porcine Biomedical Models

In translational medical research, porcine models have steadily become more popular. Considering the high value of individual animals, particularly of genetically modified pig models, and the often-limited number of available animals of these models, establishment of (biobank) collections of adequately processed tissue samples suited for a broad spectrum of subsequent analyses methods, including analyses not specified at the time point of sampling, represent meaningful approaches to take full advantage of the translational value of the model. With respect to the peculiarities of porcine anatomy, comprehensive guidelines have recently been established for standardized generation of representative, high-quality samples from different porcine organs and tissues. These guidelines are essential prerequisites for the reproducibility of results and their comparability between different studies and investigators. The recording of basic data, such as organ weights and volumes, the determination of the sampling locations and of the numbers of tissue samples to be generated, as well as their orientation, size, processing and trimming directions, are relevant factors determining the generalizability and usability of the specimen for molecular, qualitative, and quantitative morphological analyses. Here, an illustrative, practical, step-by-step demonstration of the most important techniques for generation of representative, multi-purpose biobank specimen from porcine tissues is presented. The methods described here include determination of organ/tissue volumes and densities, the application of a volume-weighted systematic random sampling procedure for parenchymal organs by point-counting, determination of the extent of tissue shrinkage related to histological embedding of samples, and generation of randomly oriented samples for quantitative stereological analyses, such as isotropic uniform random (IUR) sections generated by the &#34;Orientator&#34; and &#34;Isector&#34; methods, and vertical uniform random (VUR) sections.

The practical application and performance of methods for generation of representative tissue samples of porcine animal models for a broad spectrum of downstream analyses in biobank projects are demonstrated, including volumetry, systematic random sampling, and differential processing of tissue samples for qualitative and quantitative morphologic and molecular analyses types.

In translational medical research, porcine models have steadily become more popular. Considering the high value of individual animals, particularly of ...

Subclavian Vein Puncture As an Alternative Method of Blood Sample Collection in Rats

Blood collection with enough blood volume is essential in animal experiments. Blood collection from the tail vein of rats is popular and less stressful compared to other more aggressive methods such as retro-orbital plexus sample collection. However, this blood collection method is sometimes limited by an unsatisfactory success rate. Here, we introduce a method for blood collection through the subclavian vein puncture. The subclavian vein is located just under the clavicle and this vein is large enough to fulfill the volume requirements of blood collection. Our results show that this method is safe and applicable for blood collection sampling with the required blood volume. Blood collection through the subclavian vein puncture could serve as an alternative blood collection method in case of failed tail vein blood sampling in rats.

Here, we present a protocol to collect blood samples from the rat subclavian vein.

Blood collection with enough blood volume is essential in animal experiments. Blood collection from the tail vein of rats is popular and less stressful ...

Blood Collection Through Subclavian Vein Puncture in Mice

The mouse is the foremost mammalian model for studying human disease and human health. However, blood sample collection from mice is challenging in research work. Tail blood collection is a popular method when a small amount of blood sample is needed. Orbital artery could be considered if a large amount of blood is needed but this blood collection method has ethical issues. Formerly, we demonstrated the feasibility and safety of blood sample collection through subclavian vein puncture in rats, and here we investigate whether this method could be used in mice. We report that this method is safe and practical for blood collection in mice. Blood collection through the subclavian vein puncture in mice can be a convenient method in daily research works.

Here, we present a protocol to take blood samples from the subclavian vein of mice.

The mouse is the foremost mammalian model for studying human disease and human health. However, blood sample collection from mice is challenging in ...

Microsurgical Skills of Establishing Permanent Jugular Vein Cannulation in Rats for Serial Blood Sampling of Orally Administered Drug

Blood sampling in small laboratory animals is necessary for pharmaceutical lead optimization but can cause great harm and stress to experimental animals, which could potentially affect results. The jugular vein cannulation (JVC) in rats is a widely used model for repeated blood collection but requires adequate training of surgery skills and animal care. This article details the microsurgical procedures for establishing and maintaining a permanent JVC rat model with specific focus on the placement and sealing of the jugular cannula. The importance of monitoring physiological (e.g., body weight, food, and water intake) and hematological parameters, was highlighted with results presented for 6 days post-surgery during the rat&#39;s recovery. The drug-plasma concentration-time profile of orally administered natural phenol ellagic acid was determined in the JVC rat model.

Detailed microsurgical techniques are demonstrated to establish a longer-term jugular vein cannulation rat model for sequential blood collection in the same animal. Physiological and hematological parameters have been monitored during the rat's recovery phase. This model has been applied to study pharmacokinetics of orally administered polyphenol without inducing animal stress.

Blood sampling in small laboratory animals is necessary for pharmaceutical lead optimization but can cause great harm and stress to experimental animals, ...

Long-Term Continuous Measurement of Renal Blood Flow in Conscious Rats

The kidneys play a crucial role in maintaining the homeostasis of body fluids. The regulation of renal blood flow (RBF) is essential to the vital functions of filtration and metabolism in kidney function. Many acute studies have been carried out in anesthetized animals to measure RBF under various conditions to determine mechanisms responsible for the regulation of kidney perfusion. However, for technical reasons, it has not been possible to measure RBF continuously (24 h/day) in unrestrained unanesthetized rats over prolonged periods. These methods allow the continuous determination of RBF over many weeks while also simultaneously recording blood pressure (BP) with implanted catheters (fluid-filled or by telemetry). RBF monitoring is carried out with rats placed in a circular servo-controlled rat cage that enables the unrestrained movement of the rat throughout the study. At the same time, the tangling of cables from the flow probe and arterial catheters is prevented. Rats are first instrumented with an ultrasonic flow probe placement on the left renal artery and an arterial catheter implanted in the right femoral artery. These are routed subcutaneously to the nape of the neck, and connected to the flowmeter and pressure transducer, respectively, to measure RBF and BP. Following surgical implantation, rats are immediately placed in the cage to recover for at least one week and stabilize the ultrasonic probe recordings. Urine collection is also feasible in this system. The surgical and post-surgical procedures for continuous monitoring are demonstrated in this protocol.

The present protocol describes a long-term continuous measurement of renal blood flow in conscious rats and simultaneously recording blood pressure with implanted catheters (fluid-filled or by telemetry).

The kidneys play a crucial role in maintaining the homeostasis of body fluids. The regulation of renal blood flow (RBF) is essential to the vital ...

Comparative Study of Blood Collection Techniques in Rats

Modified Tail Vein and Penile Vein Puncture for Blood Sampling in the Rat Model

Blood samples are required in most experimental animal designs to assess various hematological parameters. This paper presents two procedures for blood collection in rats: the lateral tail vein puncture and the dorsal penile vein puncture, which offer significant advantages over other previously described techniques. This study shows that these two procedures allow for fast sampling (under 10 min) and yield sufficient blood volumes for most assays (202 &#956;L &#177; 67.7 &#956;L). The dorsal penile vein puncture must be done under anesthesia, whereas the lateral tail vein puncture can be done on a conscious, restrained animal.
Alternating these two techniques, therefore, enables blood draw in any situation. While it is always recommended for an operator to be assisted during a procedure to ensure animal welfare, these techniques require only a single operator, unlike most blood sampling methods that require two. Moreover, whereas these previously described methods (e.g., jugular stick, subclavian vein blood draw) require extensive prior training to avoid harm to or death of the animal, tail vein and dorsal penile vein puncture are rarely fatal. For all these reasons, and according to the context (e.g., for studies including male rats, during the perioperative or immediate postoperative period, for animals with thin tail veins), both techniques can be used alternately to enable repeated blood draws.

Author Spotlight: Context Dependent Blood Collection Methods in Rats 

Here, we present a protocol to offer rapid, easy, and reliable blood collection alternatives for the rat model. We describe three different blood sampling methods according to the context: tail vein puncture under anesthesia or on a conscious animal, and dorsal penile vein puncture under anesthesia.

Blood samples are required in most experimental animal designs to assess various hematological parameters. This paper presents two procedures for blood ...