This study was supported by Cuiying Plan Project of Lanzhou University Second Hospital (Grant No. PR0121015) and Gansu Provincial Key Laboratory of Urinary System Disease Research (Grant No. 0412D2).

This study adhered to the guidelines outlined in the 8th edition of the Guide for the Care and Use of Laboratory Animals13. The research received approval from the Ethics Committee of Lanzhou University Second Hospital and was documented in adherence to The ARRIVE guidelines 2.014. Twelve healthy Wistar rats (six males weighing 290-330 g and six females weighing 250-280 g) aged 12-16 weeks were accommodated in the GLP Animal Laboratory of Lanzhou University for 3 days before the actual experiment. The rat cages utilized were of the R5 type, measuring 545 mm x 395 mm x 200 mm, and were equipped with autoclaved bedding material. All rats were provided unrestricted access to both food and water. The laboratory maintained an average humidity of 25%, an average temperature of 24 &#176;C, and a light cycle that alternated between day and night (7:00 AM/7:00 PM). At the conclusion of the study, all animals were humanely euthanized using an overdose of isoflurane. For comprehensive information regarding the materials and instruments employed in this study, please refer to the Table of Materials.
1. Sample size calculation and animal selection
<ol>
	<li>Choose the resource equation method15 to estimate the animal sample size using equation (1). 
	E = Total number of animals &#8722; Total number of groups&#160;&#160;(1) 
	Where E is the degree of freedom of analysis of variance (ANOVA) and ranges from 10 to 20. 
	NOTE: In this study, 12 animals were divided into two groups A and B (three males and three females per group).</li>
	<li>Define the primary outcome of this study as the success rate and time consumption of repeated blood sampling by a single individual.</li>
	<li>Define the secondary outcome measures as changes in rat body weight, food and water intake, as well as the incidence of adverse events (such as clavicle fractures, subcutaneous hematomas, pneumothorax, and mortality).</li>
	<li>Define successful blood sampling as meeting the following criteria: i) fewer than three punctures for a single blood collection; ii) a total time (from rat restraint to completion of blood collection) not exceeding 5 min; and iii) achieving the targeted blood volume while obtaining clear plasma. Consider any deviation from these criteria a sampling failure.</li>
</ol>
2. Animal restraint and blood collection
NOTE: Blood samples from group A and B rats were collected by two experienced researchers, both of whom had drawn at least 100 blood samples. Blood samples were gathered from both groups of rats for a total of 96 times over the course of 4 days. This blood collection method does not require anesthesia or additional restraint devices for the rats. However, it necessitates adept handling techniques.
<ol>
	<li>At 8:00 AM on the day before blood sampling (day 1), assign each rat to its individual cage while its food and water are weighed. Then, have another researcher, blinded to the measurements, record the rats&#39; weight, food consumption, and water intake every day at 8:00 AM from day 1 onwards.</li>
	<li>To follow this protocol, first draw blood at 10:00 AM and then at 10:00 PM each day, collecting 0.15 mL of blood alternately from the subclavian veins on both sides. 
	NOTE: The amount of blood to be collected was determined by the maximum volume that the lowest-weight rat could tolerate within a week.</li>
	<li>Flush a syringe with sodium heparin (25 U/mL) and disinfect the injection site with alcohol.</li>
	<li>Gently stroke the rat&#39;s back skin and pinch its neck repeatedly to help the rat relax (Video 1).</li>
	<li>Using the non-dominant hand&#39;s thumb and index finger, grip and lift the rat&#39;s neck skin firmly (Figure 1A and Video 1).</li>
	<li>With coordination from the dominant hand, use the remaining three fingers and palm of the non-dominant hand to secure the rat&#39;s back skin and immobilize its front limbs (Figure 1B,C and Video 2). 
	NOTE: If the rat resists or struggles, this procedure can be repeated several times to help the rat become accustomed to the handling. The following steps are key to successful blood collection.</li>
	<li>Using the non-dominant hand&#39;s index finger, gently push down on the rat&#39;s head skin while the other fingers, along with the palm, assist in outwardly rotating the shoulder joint. During this process, use the dominant hand to fully extend the rat&#39;s shoulder joint (Figure 1D-F and Video 2).</li>
	<li>Grip the rat firmly with the non-dominant hand to align the rat&#39;s head and body in a straight line (Figure 1G,H). Then, use the dominant hand to locate the position of the clavicle and confirm the puncture site (Figure 1I, Video 2, and Video 3). 
	NOTE: Shaving the rat is not necessary. In Figure 1, shaving was done only to show the clavicle and puncture position to a greater extent. When restraining rats, especially rats &#62;350 g, allowing the rat to rest its feet on a solid surface will help support their body weight. Additionally, the restrainer should monitor the respiratory rate of each rat while collecting blood to ensure the restraint is not too tight, which may cause respiratory distress.</li>
	<li>Holding the syringe parallel to the rat&#39;s body in the dominant hand, with the needle tip facing upwards and the syringe scale toward the experimenter, maintain an approximately 15&#176; angle with the midline of the rat&#39;s body. Insert the needle 0.5 cm below the clavicle notch (at the junction of the proximal third of the clavicle and the sternum), ensuring that the needle remains parallel to the rat&#39;s body (Figure 1J and Video 3). 
	NOTE: Particular attention should be paid to the angle and depth of the needle insertion to avoid piercing the blood vessel or causing inadvertent damage to adjacent vessels.</li>
	<li>Gently withdraw the syringe slightly to create a negative pressure, often accompanied by a palpable sensation of breakthrough upon entering the blood vessel (particularly pronounced during the initial blood collection). Maintain this position and collect 0.1-1.0 mL of blood at a constant speed as needed (following the IACUC guidelines of approximately 4-5.3 mL/kg of blood per week1) (Figure 1K and Video 3).</li>
	<li>If there is no blood upon puncture, try gently adjusting the angle and depth of the needle or gently rotate the syringe (Video 3). If three consecutive attempts on the same side are unsuccessful, stop all bleeding and then switch to the opposite side for the puncture. 
	&#8203;NOTE: Swift puncture through the skin is advisable to prevent the rat from struggling due to discomfort.</li>
	<li>Apply a cotton swab for hemostasis and return the rat to its cage (Video 4).</li>
	<li>Process the blood samples according to the experimental requirements.</li>
</ol>
3. Blood sample processing
<ol>
	<li>Dispose of the syringe needle into a sharps container. Transfer the collected blood into a 1.5 mL microcentrifuge tube previously rinsed with heparin. Place the tube into a centrifuge, setting it at 4 &#176;C and 1,200 x g, and centrifuge for 10 min to separate plasma. Transfer the serum using a 1.0 mL Pasteur pipette into a clean microcentrifuge tube and store it at -80 &#176;C. 
	&#8203;NOTE: To prevent hemolysis due to pressure, remove the needle tip when necessary. During plasma aspiration, avoid drawing blood cells from the bottom of the tube. Occasionally, the surface of the syringe might collect rat fur; take care not to allow any fur to enter the tube, as it can lead to clotting.</li>
</ol>
4. Statistical analysis
<ol>
	<li>Present all data as mean &#177; standard deviation and test them for homogeneity of variance.</li>
	<li>Use Fisher&#39;s exact test to compare the success rates between groups.</li>
	<li>Use a two-sample independent t-test to compare the overall means between the two groups.</li>
	<li>Use analysis of variance (ANOVA) for continuous measurements such as blood sampling time, body weight, food intake, and water consumption. Consider P &#60; 0.05 statistically significant.</li>
</ol>

Subclavian Vein Puncture Method for Repeated Blood Collection in Conscious Rats

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(196), e65513 (2023).</li><li>Wang, L., 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=Repetitive+blood+sampling+from+the+subclavian+vein+of+conscious+rat.">Repetitive blood sampling from the subclavian vein of conscious rat.</a> Journal of Visualized Experiments. (180), e63439 (2022).</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>Tochitani, T., 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=Effects+of+microsampling+on+toxicity+assessment+of+hematotoxic+compounds+in+a+general+toxicity+study+in+rats.">Effects of microsampling on toxicity assessment of hematotoxic compounds in a general toxicity study in rats.</a> The Journal of Toxicological Sciences. 47 (7), 269-276 (2022).</li><li>Harikrishnan, V. S., Hansen, A. K., Abelson, K. S., S&#248;rensen, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comparison+of+various+methods+of+blood+sampling+in+mice+and+rats:+Effects+on+animal+welfare.">A comparison of various methods of blood sampling in mice and rats: Effects on animal welfare.</a> Laboratory Animals. 52 (3), 253-264 (2018).</li><li>Yokoyama, 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=Lack+of+toxicological+influences+by+microsampling+(50+&#181;L)+from+jugular+vein+of+rats+in+a+collaborative+28-day+study.">Lack of toxicological influences by microsampling (50 &#181;L) from jugular vein of rats in a collaborative 28-day study.</a> The Journal of Toxicological Sciences. 45 (6), 319-325 (2020).</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>Lu, 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=Microsurgical+skills+of+establishing+permanent+jugular+vein+cannulation+in+rats+for+serial+blood+sampling+of+orally+administered+drug.">Microsurgical skills of establishing permanent jugular vein cannulation in rats for serial blood sampling of orally administered drug.</a> Journal of Visualized Experiments. (178), e63167 (2021).</li><li>National Research Council of the National Academies, Committee for the Update of the Guide for the Care and Use of Laboratory Animals. <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. , (2011).</li><li>Perciedu Sert, N., 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=The+ARRIVE+guidelines+2.0:+updated+guidelines+for+reporting+animal+research.">The ARRIVE guidelines 2.0: updated guidelines for reporting animal research.</a> The Journal of Physiology. 598 (18), 3793-3801 (2020).</li><li>Charan, J., Kantharia, N. 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The authors have no relevant financial or non-financial interests to disclose.

Although tail vein blood collection is the most common method for repeated blood sampling in rats, it might be influenced by anesthesia drugs, and due to the small size of the tail vein, the amount of blood that can be collected in a single instance is limited, leading to a longer blood collection duration4,5. Although high-performance liquid chromatography (HPLC) -tandem mass spectrometry (MS/MS) systems combined with capillary microsampling (CMS) of rat tail ve...

<a href="https://app.jove.com/t/6595">This corrects</a> the article <a href="https://dx.doi.org/10.3791/66075">10.3791/66075</a>

An erratum was issued for: <a href="https://www.jove.com/t/66075">Subclavian Vein Blood Sampling in Conscious Rats</a>. The Discussion section was&#160;updated.

Erratum: Subclavian Vein Blood Sampling in Conscious Rats

Rats are one of the most common experimental animals, and there are many ways to obtain blood samples. For experiments involving a single blood collection at the concluding stage, a sufficient amount of blood can be obtained through cardiac puncture or abdominal aorta blood collection1. However, some studies require repeated blood collection from rats for routine blood or biochemical analysis, especially in pharmacokinetics and toxicology studies, where repeated blood collection is required to determine the absorption, distribution, and metabolism of drugs2.
Currently, although tail vein blood collection is the most common method for blood sampling from rats, despite not requiring anesthesia, this method can be challenging for repeated collections, and the volume of blood collected is relatively small3,4. Additionally, although blood can be collected from the saphenous and penile veins, the amount of blood obtained is limited, and anesthesia is required1,5. Furthermore, blood samples collected from the submandibular venous plexus, as well as sublingual, jugular, and subclavian veins provide higher-quality samples but typically require anesthesia or the assistance of multiple individuals1,6,7,8,9. Finally, retro-orbital sinus/canal blood collection not only necessitates anesthesia but may also potentially cause injury and stress to the rats9.
The quality of blood samples typically obtained from major veins is generally of the highest standard1. At present, some studies have found that continuous microsampling through the jugular vein is a very suitable method for toxicological research in rats although this method usually requires jugular vein catheterization10,11,12. Therefore, it is worth exploring how to obtain high-quality blood samples in accordance with the 3R principle of animal research without surgical intervention. The objective of this study was to present a method for efficiently extracting blood from the subclavian vein in rats. This technique enables the swift collection of satisfactory samples through a single-person procedure without the need for anesthesia.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.75% normal saline</td>
			<td>Gansu Fuzheng Pharmaceutical Technology Co., Ltd.</td>
			<td>&#8212;&#8212;</td>
			<td>Prepared heparin sodium solution</td>
		</tr>
		<tr>
			<td>1 mL Pasteur pipette&#160;</td>
			<td>Biosharp</td>
			<td>BS-XG-01-NS</td>
			<td>Blood collection</td>
		</tr>
		<tr>
			<td>1 mL syringe (26 G, 0.45 mm x 12 mm)</td>
			<td>Shinva Medical Instrument Co.,Ltd.</td>
			<td>0.45*12RWLB</td>
			<td>Blood collection&#160;</td>
		</tr>
		<tr>
			<td>1.5 mL Eppendorf tube</td>
			<td>Biosharp</td>
			<td>BS-15-M</td>
			<td>Blood storage and collection</td>
		</tr>
		<tr>
			<td>75% medical alcohol</td>
			<td>Shandong Lircon Medical Technology Co., Ltd.</td>
			<td>&#8212;&#8212;</td>
			<td>Disinfection of rat blood collection site</td>
		</tr>
		<tr>
			<td>Centrifuge tube holder</td>
			<td>Biosharp</td>
			<td>BS-05/15-SM60</td>
			<td>&#8212;&#8212;</td>
		</tr>
		<tr>
			<td>Electronic scale</td>
			<td>Shanghai PUCHUN Measure Instrument Co., Ltd.</td>
			<td>JE1002</td>
			<td>Weigh</td>
		</tr>
		<tr>
			<td>Heparin sodium injection</td>
			<td>Hebei Changshan Biochemical Pharmaceutical Co., Ltd.</td>
			<td>&#8212;&#8212;</td>
			<td>Rinse the syringe and EP tube; dilute with normal saline to 25 U/mL</td>
		</tr>
		<tr>
			<td>Low temperature centrifuge</td>
			<td>HuNan Xiang Yi Centrifuge Instrument&#160; Co., Ltd.</td>
			<td>&#160;H1750R</td>
			<td>Separation of serum</td>
		</tr>
	</tbody>
</table>

subclavian vein blood sampling in conscious rats

There are several established methods for obtaining repeated blood samples from rats, with the most commonly employed methods being lateral tail vein sampling without anesthesia and jugular vein sampling with anesthesia. However, most of these methods require assistance and anesthetic equipment and sometimes pose difficulties in terms of blood collection or the poor quality of blood samples. In addition, these methods of blood collection consume significant time and human resources when repeated blood sampling is required for a large number of rats. This study presents a technique for repetitive blood sampling in non-anesthetized rats by a single proficient individual. Highly satisfactory blood samples can be obtained by puncturing the subclavian vein. The method demonstrated an impressive overall success rate of 95%, with a median time of merely 2 min from rat restraint to the completion of blood collection. Furthermore, performing consecutive blood collections within the designated range does not inflict any harm on the rats. This method is worth promoting for blood collection, especially in large-scale pharmacokinetic studies.

Author Spotlight: Developing an Efficient Blood Collection Method in Non-Anesthetized Rats 

Subclavian Vein Blood Sampling in Conscious Rats

The scope of our research is centered on devising infection methodologies for collecting blood samples from rats. Our aim is to ensure rapid and set procedures that align with our principle, utilizing minimal manpower and resources. One of the current experimental challenges is to effectively immobilize the rat in a manner that simultaneously maintains the subclavian vein enclosed proximately to the skin surface for optimal blood collection.The advantages of our protocol is that high quality blood samples can be obtained quickly by a person without anesthesizing the rats.

High-quality plasma specimens exhibit a pale-yellow hue, clarity, and transparency, devoid of any red tinge or clotting, as depicted in Figure 2A. Figure 2B shows hemolysis (left side) or coagulation (right side) as a result of improper procedures, respectively. Over the course of 96 blood collection sessions within 4 days, the average single blood collection times for groups A and B were 119.87 &#177; 33.62 s and 123.28 &#177; ...

Watch this Scientific Journal Video about Developing an Efficient Blood Collection Method in Non-Anesthetized Rats at JoVE.com

Here we present a combination of effective rat restriction and subclavian vein puncture methods that enable rapid, safe, and repeated blood collection in rats without anesthesia.

There are several established methods for obtaining repeated blood samples from rats, with the most commonly employed methods being lateral tail vein ...

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1.2K Views.  Lanzhou University Second Hospital. The scope of our research is centered on devising infection methodologies for collecting blood samples from rats. Our aim is to ensure rapid and set procedures that align with our principle, utilizing minimal manpower and resources. One of the current experimental challenges is to effectively immobilize the rat in a manner that simultaneously maintains the subclavian vein enclosed proximately to the skin surface for optimal blood collection.The advantages of our protocol is that high q...

Video: Author Spotlight: Developing an Efficient Blood Collection Method in Non-Anesthetized Rats 

Subclavian Vein Puncture Blood Collection and Observation of Secondary Outcome Measures in Rats

On the day of the blood sampling, place the rat in the cage at 8:00 AM.Weigh the food and water and place them in the cage. Define successful blood sampling based on the following criteria. Have a blinded researcher record the rat's weight, food consumption, and water intake daily at 8:00 AM.Next, gently stroke the rat's back skin and pinch its neck repeatedly to help the rat relax.Using the non-dominant hand's thumb and index finger, grip and lift the rat's neck skin. Using the remaining three fingers and the palm of the non-dominant hand, secure the rat's back skin, and with the help of the dominant hand, immobilize its front limbs. Using the non-dominant hand's index finger, gently push down on the rat's head skin.Rotate the shoulder joint outwardly with the other fingers along the palm while the dominant hand fully extends the shoulder joint. Grip the rat firmly with the non-dominant hand to align its head and body in a straight line. Locate the clavicle with the dominant hand and confirm the injection site.Disinfect the site with alcohol, then flush the syringe with sodium heparin. Hold the syringe parallel to the rat's body and the dominant hand with the needle tip facing upwards and the syringe scale toward the experimenter. Insert the needle 0.5 centimeters below the clavicle notch.Gently withdraw the syringe to create negative pressure, collecting 0.1 to 1 milliliters of blood at a constant speed. Apply a cotton swab for hemostasis and return the rat to its cage. Average blood collection times for groups A and B were similar over 96 hours.Blood collection success rates were 93.8%for group A and 95.8%for group B, with no significant difference. Group A had a shorter average blood collection time on the second day compared to the fourth day. The number of attempts for successful blood collection averaged 1.21 for group A and 1.17 for group B.More attempts and longer puncture times indicated higher failure rates.Both groups showed steady weight gain over four days. Water and food intake remained relatively constant among rats of the same sex.