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

Podsumowanie

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.

Streszczenie

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's recovery. The drug-plasma concentration-time profile of orally administered natural phenol ellagic acid was determined in the JVC rat model.

Wprowadzenie

Repeated acquisition of blood samples from small laboratory animals, such as rodents, guinea pigs, and rabbits, is an important aspect for pharmaceutical lead optimization and also for reducing the number of animals used in research^1,2. The pipeline for developing new diagnostic tools and drug formulation (e.g., vaccine) requires access to different volumes of blood in order to evaluate their robustness and performance in vivo, such as pharmacokinetics (PK), toxicity, and sensitivity^3,4,5.

The laboratory approach to blood sample collection is broadly classified into two types, surgical and nonsurgical⁶. The nonsurgical approach is relatively easy to grasp for the researcher, which includes common techniques, such as cardiac puncture, orbital sinus puncture, and bleeding of the saphenous and tail vein. Multiple blood sampling is possible by some non-surgical methods, but the sample volume is small and can cause physical wound and psychological stress to the animals¹. On the other hand, the surgical approach is a favorite alternative to repeated venipuncture, and it involves placement of a temporary or permanent cannula in the blood vessels of animals^7,8,9. The large blood volume could be repeatedly withdrawn through the cannula in conscious rats while avoiding the stress and pain due to the handling technique, restrain, and anesthesia^7,8,10,11. However, the cannula implantation requires an experienced researcher with adequate training in order to successfully collect the blood.

Blood collection through jugular vein cannulation (JVC) in rats is the most widely used method to study the drug PK^6,10,12,13. Yet, establishment of the JVC rat model needs careful practice of microsurgical skills and knowledge of postsurgical care and maintenance. Especially, after the surgery, the rat requires administration of analgesics and sufficient recovery time to reach stable physiological condition for further experiments^13,14,15. Although the body weight gain (i.e., >10 g) is a valid and commonly applied indicator for the rat's recovery, it is not uncommon that the rats have unexpected death postoperatively due to dehydration, infection, and inflammation, which could be subtle to notice at the early onset^14,15. In addition, catheter obstruction in the JVC model remains to be an issue during the blood collection.

The present protocol has demonstrated in detail the microsurgical procedures for JVC in an anesthetized rat with specific focus on the identification, isolation, and cannulation of the jugular vein. The importance of physiological and hematological monitoring of the rats during the recovery phase is highlighted. Finally, serial blood samples were collected through the venous catheter to study the PK of the orally administered natural phenol ellagic acid with poor bioavailability (i.e., low systemic concentration) to verify the JVC rat model.

Protokół

The procedures described below were performed as part of a protocol approved by the Institutional Animal Care and Use Committee of Northwestern Polytechnical University (No. 202101117).

1. Preoperative preparation (the day before the surgery)

NOTE: Required solutions: normal saline (0.9% w/v sodium chloride), heparinized saline (1% w/v heparin sodium), catheter lock solution, non-steroidal anti-inflammatory drug (NSAID), such as meloxicam solution (2 mg/mL).

1. Solution preparation
   1. Aliquot 200 µL of pre-packaged catheter lock solution in a 1.5 mL sterile microcentrifuge tube.
      NOTE: Catheter lock solution is composed of heparinized saline (0.4% v/v heparin sodium) mixed with glycerol (v/v,1:1).
   2. Mix 1 g of heparin sodium in 100 mL of the normal saline to prepare 1% heparinized saline.
   3. Dissolve meloxicam in normal saline to prepare a 2 mg/mL concentration solution for postoperative pain relief.
      NOTE: Prepared heparinized saline and meloxicam solution are filtered through a 0.22 µm filter. All the solutions are sterilized and stored at 4°C for future use.
2. Surgical instruments and materials
   1. Pack all clean surgical tools in a pouch and tape it with a piece of autoclave sterilization tape. Refer to Figure 1A for the specific surgical instruments used.
   2. Autoclave the surgical pouch at 121 °C for 30 min for the next day use.
3. Animal preparation
   1. Prior to the surgery, house all male Sprague-Dawley (SD) rats in the standard Animal Room with controlled temperature at 22 ± 1 °C. Feed them with the standard laboratory food and water ad libitum for at least 7 days.
      NOTE: Both male and female rats can be used for the JVC model, and their typical ages and body weights vary from 9-14 weeks and 294 ± 57 g, respectively.
   2. Anesthetize the rat with 3%-3.5% isoflurane mixed with oxygen in a pre-anesthesia chamber. Determine whether the rat becomes unconscious by the lack of response to foot pinch.
   3. Gently take the rat out, place the rat's nose into an anesthetic nosepiece supplying 2%-2.5% isoflurane.
   4. In the ventral and dorsal position, remove the fur thoroughly around its right shoulder and posterior areas of neck with depilatory cream and a pet razor. Return the rat to the cage for surgery to be performed on the next day.

2. Before the surgery on the day

1. Prepare the aseptic workstation
   1. Spray 75% medical alcohol to disinfect the operation area, and then place the heating pad covered with a clean cushion. Set the LED lamp with a cold light source beside the workstation.
   2. Pre-warm the required solutions (step 1.1) to room temperature.
   3. Fill 0.6 mL of heparinized saline and 0.15 mL of catheter lock solution in two sterile 1.0 mL blunt tipped syringes, respectively. Withdraw 2.5 mL of the normal saline using a sterile 5.0 mL syringe.
   4. Soak the cotton balls in 75% medical alcohol. Squeeze out excess ethanol before use.
   5. Weigh and record the rat's body weight.

3. During the surgery

1. Surgical preparation
   1. Wear the surgical coat, sterile gloves, and facial mask. Then open the sterilized surgical pouch, leave all surgical tools in 75% medical alcohol, and dry them before use.
2. Jugular vein isolation
   ​NOTE: The estimated operation time for this part is 10 min.
   1. Anesthetize the surgery-ready and shaved rat with 3%-3.5% isoflurane mixed with oxygen in an induction chamber and determine whether the rat becomes unconscious by the lack of response to foot pinch.
   2. Place the rat's nose into the nosepiece supplied with 2%-2.5% isoflurane to maintain the anesthesia.
   3. Subcutaneously inject (s.q.) meloxicam solution at a dose of 2 mg/kg. 
      NOTE: Make sure to select analgesics that do not interact with the drug compound of interest in the pharmacokinetics study.
   4. Using adhesive tape, restrain the rat's forearms in their ventral position to each side of the surgical platform.
   5. Gently scrub the surgical area by alternating between cotton balls soaked in 75% medical alcohol and iodine-based scrub for a total of three times.
   6. Carefully lift the skin near the clavicle on the right side of the midline of the neck with forceps and make an incision towards the chest about 1.5-2.0 cm in length with a pair of surgical scissors.
   7. Blunt dissect the thin tissue cover with iris scissors to expose the underneath jugular vein. The proximal cephalic end of the external jugular vein consists of two branches, which can be visually identified.
      NOTE: Depending on the age and sex of the rat, the soft tissue (e.g., salivary glands, lymphatic nodes, and fatty tissues) covering the jugular vein varies. Compared to the young rats, the old rats are fatter (e.g., BW > 300 g), and thus need more tissue separation before the jugular vein is visible.
   8. Lift the jugular vein along with its connective membranous tissues to visualize the lymph gland attached to the jugular vein. Carefully separate the vein along the vascular direction from surrounding muscle, fat, and other tissues.
   9. Nudge the forceps under the jugular vein without damaging the collateral blood vessels and pass two pieces of 6-0 suture under the vein to mark the two ends of the blood vessel individually.
   10. Pull one piece of the suture as far as possible toward the rat head and ligate the vein cranially with 2-3 knots using forceps.
   11. Place the second ligature on the caudal end of the vein with 1 loose knot.
3. Jugular vein cannulation
   NOTE: The estimated operation time for this part is 15 min.
   1. Open the package containing 11 cm polyurethane (PU) catheter (I.D. 0.6 mm x O.D. 0.9 mm, Figure 1B) and attach the catheter to the prepared blunt tipped syringe filled with the heparinized saline.
   2. Slowly push the heparinized saline into the catheter to avoid air bubbles.
   3. Nudge the non-tip flat side of the forceps under the jugular vein to exit on the other side. Make a small v-shaped cut on the vein near the cranial tie with a pair of castroviejo micro scissors and gently open the incision with the tip of the elbow vessel dilator forceps.
      NOTE: Rinse the incision with pre-warmed normal saline (37 °C) if a small amount of blood gushes out.
   4. Cut out the oblique opening of the front end of the jugular vein catheter. Clamp the oblique end of the tube with forceps and slide it into the jugular vein.
      NOTE: This step may need another person to facilitate the catheter sliding.
   5. While advancing the catheter, slowly withdraw the elbow microsurgical forceps and clamp the outer surface of the vessel with forceps.
      NOTE: If the right blood vessel is selected and the tip of the catheter is successfully slid into the blood vessel, the entire catheter insertion process should not feel any flow resistance.
   6. Stop inserting the catheter upon hitting the first blue mark of the PU tube (Figure 1B), which is approximately 3.0 cm in length.
   7. Secure the inserted catheter to the vein with both caudal and rostral ligatures using forceps.
   8. Thread a 6-0 suture through the exposed tissue on the right side of the incision using a suture needle (1/2 curved cutting, 12 mm) and tie the ligature with a hemostat.
   9. Bend the catheter at the second blue mark (Figure 1B) to bind with the same ligature (in step 3.3.8) and to avoid occluding the PU-tubing.
   10. Snip all the extra suture thread and close the catheter by replacing the blunt tipped syringe with a 22 G stainless-steel plug.
4. Catheter exteriorization
   NOTE: The estimated operation time for this part is 10 min.
   1. Place the rat in the dorsal position and gently clean the area between the scapulae with the cotton ball soaked in 75% medical alcohol.
   2. Make a very small incision at the center of the dorsal neck with surgical scissors. Through the dorsal incision, guide and gently push the trochar underneath the skin toward the ventral incision on the right side of the neck.
   3. Put the venous catheter into the trochar and then pull out and guide the venous catheter toward the dorsal incision.
   4. Secure the exteriorized catheter into the muscle layer in the same way with the suture (see the procedure in steps 3.3.8 and 3.3.9).
   5. Close the skin layer of ventral and dorsal incisions with the 6-0 nylon suture and suture needle (3/8 curved cutting, 17 mm). Swab all surgical incisions with iodophor.
      NOTE: The wound-clips is an alternative method to close the skin incision.
   6. Remove the catheter plug by clasping the catheter with fingertips. Place a new blunt tipped syringe and slowly draw back the syringe to test the blood flow.
      NOTE: Since the rat is in the supine position, one may not be able to obtain blood samples. Blood samples could be obtained by changing to a side body position.
   7. Hold the catheter again with fingertips and inject 0.2 mL of heparinized saline and 0.1 mL of lock solution into the catheter using the blunt tipped syringe.
   8. Hold the catheter with fingertips and replace the syringe with a stainless-steel plug. Unclamp the catheter and push the plug slightly in to ensure the tightness of the catheter.

4. Immediate post-surgical care

1. Recover the rat in the dorsal decubitus position by caging it individually with fresh corncob bedding. Often, provide a temperature-regulated heating pad to maintain the core body temperature.
   NOTE: For animal welfare, leaving food and water on the bedding is an effective way to alleviate the pain caused by neck movements when eating and drinking.
2. Record the end time of the surgery and monitor the rat at 2 h intervals for at least 4 h. Provide additional analgesia for the recovery if the rat shows signs of pain or distress.

5. Physiological and hematological monitoring during recovery phase

1. Monitor the body weight and the food and water intake daily and record the data. 
2. To collect a small volume of fresh blood for hematological test, place the rat in a restrainer. Open the plug and insert the syringe into the venous PU catheter to ensure the catheter is not obstructed.
   NOTE: The blood collection was performed at the same time daily for 6 consecutive days.
3. Discard the initial withdrawn blood, which contains a mixture of blood, heparinized saline, and catheter lock solution.
4. Use a new syringe to collect 150 µL of fresh blood sample and transfer the blood sample to the 0.5 mL tube containing K2EDTA (1.8 mg/mL blood) spray dried on the tube wall.
   NOTE: If the catheter is blocked, 0.2 mL of heparinized saline can be injected into the catheter to flush the catheter a few minutes before the next blood collection time.
5. Inject sterile saline in the same volume to compensate for the withdrawn blood. Inject 150 µL of pre-warmed normal saline (37 °C) and infuse 0.2 mL sterile heparinized normal saline through the catheter.
6. Inject 100 µL of the lock solution into the catheter to ensure the sealing and sterility of the catheter before the next sample collection.
7. Analyze the blood samples within 2 h of collection using an automated blood cell counter.

6. Repeated blood sampling for pharmacokinetic studies of oral administered drug

NOTE: Rats with weight gain >10 g and stable hematological level are suggested to be enrolled for future study. Following the current protocol, the JVC rats required 4 to 6 days to recover.

1. After 4-6 days of surgery, fast the rat for 12 h with free access to water.
   NOTE: Depending on the experimental goal, fasting the animal is optional.
2. Orally gavage the fasted rat with natural phenol bioactive ellagic acid at a dose of 6 mg/kg with a straight gavage needle¹⁶.
3. Collect 200 µL of blood samples in the heparinized tubes via the jugular vein cannula at pre-determined time points over 24 h post-oral administration. The blood collection process follows the procedure in step 5.5.
   NOTE: The catheter does not need to be closed with the lock solution until the blood collection is completed.
4. Immediately centrifuge the blood sample at 3000 x g at 4 °C for 10 min.
5. Analyze the extracted plasma sample by liquid chromatography-mass spectroscopy^17,18.

Wyniki

This protocol has thoroughly demonstrated how to establish a long-term JVC model using microsurgical skills for serial blood collection. Figure 1A shows the essential surgical instruments and materials used to conduct the surgery. The specification of PU catheter with three blue marks is also illustrated, which is helpful for guiding the researcher to place the vein cannula in step 3.3., how to use the marks on the PU catheter to guide the cannulation (Figure 1B...

Dyskusje

Mastering the technique of vessel cannulation requires significant practice and learning the lesson from each operation. Christakis et al. using cumulative sum (CUSUM) analysis, found that a researcher needs to practice 200 rats over a period of one year before being ready for the PK evaluation of drug candidates²⁰. Yet, the operating time required for the vein cannulation can be significantly reduced by the number of rats performed^13,20. ...

Materiały

Name	Company	Catalog Number	Comments
0.5 mL test tube containing EDTA anticoagulant	Xinkang	N/A	collecting blood samples for hematology test
0.5*20 mm 1.0-mL syringe	KLMEDICAL	N/A	washing or replacing the fluid with saline
0.6*28.5 mm 5.0-mL syringe	HD	N/A	Subcutaneous injection
1.0-mL Blunt tipped syringe (22G)	skillsmodel	S4-PKT22G	Inject the saline and collect blood samples through catheter
1.5 mL sterile microcentrifuge tube	Axygen	MCT-150-C-S	Store sterile catheter lock solution heparinized saline and meloxicam solution
1.5 mL microcentrifuge tubes	Biosharp	BS-15-M	blood collection
1/2  circle cutting 5*12 mm suture needle	skillsmodel	S4-FHZ	Thread the muscle layer to fix the catheter
3/8 circle cutting 7*17 mm suture needle	skillsmodel	S5-FHZ	Suture the incision of rat cortex
6-0 sterile non-absorbable silk suture thread	JUNSHENG	N/A	ligature
75% medical alcohol	HONGSONG	N/A	Disinfection
Adhensive tape	LIUTAI	N/A	positioning the rat
Autoclave sterilization tape	Biosharp	BS-QT-028	Mark sterilized items
Automated blood cell counter	Sysmex	XN-550	Hematology test
Castroviejo micro scissors	skillsmodel	WA1010	Cut the opening in the blood vessel
Centrifuge	Thermo Fisher Scientific	75002402	Plasma preparation
Clean cushion	Qingjie	N/A	Prepare the operation area
Cotton balls	HC	N/A	Wound disinfection and sterilization
Cotton swabs	BEITAGOGO	N/A	Disinfection
Curved hemostat	skillsmodel	N/A	ligature
DN50 Stainless-steel rat restrainer	skillsmodel	S4-RGDQ1	Restrict the movement of rats for easy operation
Ellagic acid	Aladdin	E102710-25g	natural phenol for oral administration
Half-curved forceps	skillsmodel	53072	Lift the muscle layer and tissue, isolate the jugular vein and tie the suture
Heating pad	Warm mate	N/A	preventing heat loss of animal
Heparin sodium	Solarbio	H8060	anticoagulant
Iodophor	Xidebao	N/A	Clean the wound
Iris scissors	skillsmodel	54002	Bluent separation the muscle layer
Isoflurane	RWD	R510-22-16	anaesthesia
LED lamp	EMPERORFEEL	N/A	sugery
Liquid chromatography-mass spectroscopy	Thermo Fisher Scientific	VQF01-20001/ TSQ02-10002	detection of drug concentration in plasma
Meloxicam	Hongqiang	N/A	Analgesic
Normal saline	KL	N/A	Prepara the solution and protect blood vessels from drying out
Pet razor	Codos	3180	Shaving the fur
Phosphate-buffered saline	ZHHC	PW012	Preparation of Ellagic acid solution
PU catheter	skillsmodel	RJVC-PU	Jugular vein cannulation
Small animal operation anesthesia console	RWD	68620	Operation workstation
Spray bottle	Other	N/A	aseptic workstation
Stainless steel plug (22G)	skillsmodel	S4-PKD22G	Plug the catheter to ensure its sealing
Stainless steel trochar	skillsmodel	S$-PKDGZ	Guide the catheter exteriorization
Sterile lock solution	skillsmodel	SK-FB	lock the catheter to ensure its sterility
Straight feeding needle	skillsmodel	N/A	Oral gavage
Surgical pouch	BKMAM	N/A	container for sterilization of surgical instruments
Surgical scissors	skillsmodel	J21070	Cut incision on rat skin
Vessel dilator balanced forceps	skillsmodel	WA3020	Expand the blood vessel and guide the cannula to slide in
ZS-MV Small animal anesthesia machine	ZSLab	1057003	inducing and maintaining anaesthesia