A Single-Operator Jugular Vein Sampling Method for Repeated Blood Collection in Conscious Rats

Abstract

Jugular vein (JV) blood sampling is widely employed in pharmacological, toxicological, and pharmacokinetic studies due to its ability to provide large and repeatable blood volumes. However, current approaches are constrained by low convenience, safety concerns, and variable sample quality. Here, we report the development of a new single-operator JV blood sampling technique for conscious rats. By optimizing animal restraint, puncture site localization, and procedural standardization, the method significantly reduced sampling time, puncture attempts, pain-related behaviors, and hemorrhagic injury, while ensuring consistent blood yield. Compared with the conventional approach, it achieved a 100% success rate, halved the average sampling time, and maintained stable serum biochemical, coagulation, and hematologic parameters. The utility of this method was further confirmed in an intraperitoneal glucose tolerance test and a cisplatin-induced acute kidney injury model, where data consistency was preserved. This technique provides a convenient, safe, and reliable alternative for repeated blood collection in conscious rats, ensuring animal welfare and supporting the generation of high-quality data in preclinical research.

Introduction

Rats are widely used in pharmacological, toxicological, and pharmacokinetic studies, in which blood samples collected at defined time points are essential for evaluating drug efficacy and metabolic profiles. The choice of sampling technique directly influences both sampling efficiency and blood quality, which in turn determine the reliability of experimental outcomes. For single terminal collections, methods such as cardiac puncture or abdominal aorta sampling can yield sufficient volumes of blood.^1,2 However, these approaches are terminal procedures and thus unsuitable for repeated sampling. In contrast, pharmacokinetic and toxicological investigations typically require multiple sequential blood collections to assess biochemical parameters and elucidate absorption, distribution, and metabolic processes.³ Therefore, the development of efficient, safe, and reproducible techniques for repeated blood sampling in rats is of critical importance.

A variety of blood sampling methods from various vessels are currently available for rats, including the tail vein, jugular vein (JV), retro-orbital venous plexus, suborbital sinus, submandibular vein, saphenous vein, and penile vein. Each approach has specific advantages but also notable drawbacks. Tail vein blood collection is a commonly used and relatively safe method, but it yields only small volumes and is time consuming; although warming devices can accelerate the procedure, they require additional equipment and may compromise sample quality, thereby limiting its applicability in studies requiring blood collection at defined time points.⁴ Retro-orbital plexus sampling is technically convenient but carries a risk of ocular injury and vision impairment,⁵ while suborbital sinus puncture requires anesthesia, increasing the risk of tissue trauma and stress.⁶ Submandibular vein sampling is rapid and minimally invasive, yet it is associated with higher complication rates and inconsistent sample quality.⁷ Sampling from the saphenous or penile vein is restricted by the small obtainable volume.⁸ To overcome these limitations, anesthesia or auxiliary devices are often employed to facilitate repeated collections; however, such measures increase experimental costs, reduce efficiency, and may compromise sample integrity or alter metabolic homeostasis in rats.⁹ These challenges highlight the need for alternative blood sampling strategies that are specifically suited for repeated applications in toxicological and pharmacological studies.

Recent advances in venous blood collection techniques have emphasized JV access as a high standard for blood collection because of the superior sample quality obtained. Permanent JV catheterization allows long-term serial microsampling,^10,11 but its requirement for surgical implantation limits practical application.¹² Subclavian vein puncture under anesthesia can yield sufficient volumes due to the vessel’s large caliber, but it depends on systemic anesthesia and carries substantial procedural risks.¹³ Later adaptations, such as vertical subclavian vein puncture in conscious rats, eliminated anesthesia but required 2 operators for effective restraint and posed considerable risk of hemorrhage.¹⁴ More recently, a conscious subclavian vein sampling technique has been described, offering advantages of single-operator feasibility, high convenience, and low cost.¹⁵ However, the safety and hematologic stability associated with this method remain insufficiently validated, and the close anatomic relationship of the subclavian vein with thoracic structures and brachial plexus nerves introduces risks such as pneumothorax, hematoma, and nerve injury. Thus, current jugular and subclavian vein methods still face notable limitations in convenience, safety, and sample quality, highlighting the need for techniques that ensure efficient, safe, and stable repeated blood collection in conscious rats.

To address these gaps, we developed a new single-operator, repeated blood sampling technique using the JV in conscious rats. This method employs a standardized procedure to achieve precise puncture site localization and alternating bilateral sampling, enabling rapid, high-quality blood collection without auxiliary assistance. Using established JV methods as controls,¹⁴ we systematically evaluated the new approach through repeated sampling experiments, behavioral assessments, histopathology, hematologic and biochemical analyses, and validation in pharmacokinetic and disease model settings. Our findings provide strong evidence that this method offers enhanced convenience, safety, and blood sample stability, supporting its application as a reliable tool in preclinical research.

Materials and Methods

Chemicals and reagents.

Tiletamine hydrochloride and zolazepam hydrochloride for injection (Zoletil 50) were purchased from Suzhou Kunchen Biotechnology (Suzhou, China). A 10% formalin solution (Type B; 10 L) was obtained from Zhongshan Kangnaixin Biomedical Technology (Zhongshan, China). Hematoxylin and eosin (H&E) staining solutions (1,000 mL/bottle) were provided by Zhuhai Beisuo Biotechnology (Guangdong, China). Anhydrous glucose (purity: 99%; lot no. KYIFJ031; CAS no. 50-99-7) was purchased from Inoke Technology (Shanghai, China). Phosphate buffered saline (PBS) was purchased from Dingguo Changsheng Biotechnology (Beijing, China). Cisplatin (CAS no. 15663-27-1) and DMSO (Molecular Biology Grade; CAS no. 67-68-5) were obtained from Sigma-Aldrich (Beijing, China). Normal saline was provided by Chifeng Yuansheng Pharmaceutical (Chifeng, China). All reagents were used within their validity period and met the required pharmaceutical grade standards.

Animals.

To minimize interference from physiologic variables in female animals, 24 male Sprague-Dawley (SD) rats (7-8 weeks old; 200-210 g) were purchased from the Guangdong Laboratory Animal Center (Guandong, China) and acclimated for at least 7 days before the study initiation. All rats were from specific pathogen-free (SPF) colonies, which excluded Salmonella spp., Mycoplasma spp., Corynebacterium kutscheri, Clostridium piliforme, Rodentibacter pneumotropicus, Klebsiella pneumoniae, Pseudomonas aeruginosa, Bordetella bronchiseptica, hantavirus, Sendai virus, pneumonia virus of mice, reovirus type 3, rat parvovirus RV strain and H-1 strain, ectoparasites, Toxoplasma gondii, flagellates, ciliates, and all helminths. Rats were placed in individual isolated cages (466 × 314 × 200 mm; Suzhou Suhang Technology Equipment, Suzhou, China) lined with corncob bedding (production license no. SCXK [Jing] 2020-0010; Ke’ao XieLi Feed, Beijing, China). The housing environment was maintained under constant conditions: 70 ± 2 °F (22 ± 2 °C), relative humidity of 30%-70%, and a 12-hour light/dark cycle (7:00 am-7:00 pm). Rodent feed (Ke’ao XieLi [Tianjin] Feed) and reverse osmosis water (CNAS, ILAC-MRA Certification; no. GZ250811040-00001; Jinzhi Testing Technology, Guangzhou, China) were provided ad libitum. The feed complied with the requirements for experimental animal feed composition specified in GB 14924.3-2010, and the drinking water met the standards of GB 5749-2022. All cages, cage components, bedding, and equipment were autoclaved before use (product specification: SGLS-A-1500D; Genuo Biotechnology, Shenzhen, China) at 121 °C for 30 minutes. During the entire study period, 4 rats were housed per cage. The study was conducted at Guangdong Sunshine Lake Pharma (license no. SYXK [Yue] 2023-0135) and approved by the Institutional Animal Ethics Committee (approval ID: IAEC-K-250605-01). Sample size determination was based on the resource equation method¹⁶ and experimental design requirements. All animal experiments were conducted in compliance with the Guide for the Care and Use of Laboratory Animals (8th ed)¹⁷ to minimize unnecessary pain and distress.

JV blood sampling techniques.

The traditional JV sampling technique (Figure 1A) was performed based on previously reported procedures.¹⁴ Briefly, (1) animal restraint: the rat was placed on the cage rack. The operator used the left thumb and forefinger to grasp the skin at the right side of the neck and back below the ear (Figure 1A-a), with the remaining fingers and palm holding the dorsal skin to keep the head and thorax in an upright position (Figure 1A-b). (2) Venipuncture procedure: facing the ventral side of the rat, an assistant extended the forelimb on the sampling side. The puncture site was located at the lowest point of the subclavian triangle between the neck and forelimb (Figure 1A-c). The operator inserted the needle at a 45° outward angle, aligned parallel to the longitudinal axis of the head, with the bevel directed toward the abdomen (Figure 1A-d). The injection needle was advanced from a shallow to a deep position, with simultaneous retraction to create negative pressure. When blood flowed into the syringe, needle insertion was stopped, the needle was stabilized, and the required volume of blood was drawn at a constant rate. (3) Hemostasis: after sampling, the needle was withdrawn, and gentle compression was applied with a sterile cotton ball for 3-5 seconds to ensure hemostasis (Figure 1A-e).

Figure 1.

Jugular Vein (JV) Blood Sampling Techniques in Rats. (A) Traditional method: (a) grasping; (b) restraint; (c) puncture site (red dashed cross), located at the lowest point of the subclavian triangle between the neck and forelimb (black arrows indicate the corresponding anatomic structures); (d) blood collection; and (e) hemostasis. (B) New method: (a) grasping; (b) restraint; (c) puncture site (red dashed cross), located at the midpoint between the most prominent point of the shoulder joint and the thoracic midline (black arrows indicate the corresponding anatomic structures); (d) blood collection; and (e) hemostasis. (C) Cervical anatomy: (a) puncture sites of the 2 methods and (b) anatomic image of the left neck and thorax in normal rats (without repeated blood collection), with partial removal of skin and muscle tissues. White box highlights the JV region, and arrows indicate the relevant structures.

The new JV sampling technique is illustrated in Figure 1B. Its main modifications involved restraint, puncture site localization, and venipuncture procedure: (1) animal restraint: the rat was positioned naturally on the cage rack. The operator gently lifted the rat’s upper body with the left hand and rested it on the right palm (Figure 1B-a). The left thumb and middle finger secured the left and right forelimbs to the dorsum in a crossed manner. Subsequently, the index finger slid from between the rat’s ears to the raised dorsal skin, then clasped tightly with the thumb and middle finger; at this point, the 3 fingers were used to secure the rat’s dorsal skin (Figure 1B-b). Simultaneously, the ring and little fingers grasped the proximal half of the tail, ensuring that the head, neck, and thorax were maintained on the same horizontal plane to avoid cervical torsion. (2) Venipuncture procedure: with the syringe bevel facing upward, the needle was inserted at an angle of approximately 20° relative to the thoracic plane. The puncture site was located at the midpoint between the most prominent point of the shoulder joint and the thoracic midline (Figure 1B-c). The needle was advanced slowly to a depth of less than 1 cm while gentle negative pressure was applied, and once blood entered the syringe, aspiration was continued steadily until the required volume was obtained (Figure 1B-d). (3) Hemostasis: the needle was withdrawn, and gentle compression was applied with a sterile cotton ball for 3-5 seconds to ensure hemostasis (Figure 1B-e).

Notably, operators were required to demonstrate proficiency in both restraint and venipuncture skills before performing the procedure independently (prior training and practice of more than 1 week are required, with successful collection of over 200 μL of blood from 50 rats). The comparative puncture sites of the traditional and new techniques are illustrated in Figure 1C-a and C-b.

Repeated JV sampling experiment.

To evaluate the practical performance of the new method, the 2 techniques were compared in terms of convenience, including sampling duration, success rate, and puncture frequency. Evaluation criteria were based on a previous¹⁵ study: (1) effectiveness indicators: sampling was deemed successful if all of the following were met: collection time less than or equal to 180 seconds, puncture attempts less than or equal to 3, and blood volume greater than or equal to 200 μL. Any failure in one criterion was recorded as unsuccessful; (2) animal welfare termination criteria: experiments were terminated, and euthanasia performed if rats exhibited greater than or equal to 20% body weight loss, clavicle fracture, hematoma greater than or equal to 1 cm in diameter, or pneumothorax (respiratory distress, cyanosis); and (3) blinding: all operators were qualified laboratory animal technicians trained in both methods and blinded to group allocation to ensure objectivity.

After a 1-week acclimatization period, the rats were randomly allocated into 3 groups (n = 8 per group) using the Excel RAND () function: Control, traditional method (TM), and new method (NM) groups. To replicate intensive sampling conditions typically encountered in pharmacological and toxicological studies,^12,15 a 7-day schedule comprising 13 sampling time points was employed, with 2 collections performed daily at 9:00 am and 5:00 pm. In the TM group, 2 operators were required, one responsible for animal restraint and the other for venipuncture, whereas in the NM group the procedure was completed by a single operator. To avoid repeated puncture at the same site and thereby minimize local tissue injury, blood was collected alternately from the left JV in the morning and the right JV in the afternoon. At each time point, 200 μL of blood were withdrawn using a 1-mL syringe fitted with a 0.3-mm needle (Jiangnan Medical Equipment, Shenzhen, China). The duration of sampling, the number of puncture attempts, and the success rate were carefully recorded. Rats in the control group did not undergo any intervention and were monitored only for body weight changes, serving as baseline controls. Upon completion of the repeated sampling protocol, all animals were returned to standard housing conditions for subsequent experimental procedures. A schematic overview of the experimental workflow is presented in Figure 2A.

Figure 2.

Assessment of Procedural Convenience in JV Blood Sampling. (A) Experimental workflow. (B) Body weight (n = 8). (C) Hemolysis in serum/plasma samples (n = 8). (D) Hemoglobin concentration in serum/plasma (n = 8). (E) Sampling time per attempt (n = 8). (F) Mean overall sampling time across 13 collections (n = 104). (G) Puncture attempts per collection (n = 8). (H) Mean overall puncture attempts across 13 collections (n = 104). (I) Success rate of each sampling session (n = 8). (J) Mean overall success rate across 13 sessions (n = 13). Data are presented as mean ± SD (n = 8); *P < 0.05, **P < 0.01, ***P < 0.001 vs. TM group; independent-samples t test.

Behavioral observation.

Behavioral assessments were conducted to further evaluate the safety of the sampling method. During repeated sampling, a senior veterinary technician (3 years of work experience) blinded to group assignments observed the animals for normal behaviors and pain- or stress-related behaviors.¹⁸ Normal behaviors included feeding or drinking, grooming, rearing on hind limbs (high posture), and standing on all 4 limbs (low posture). Pain- or stress-related behaviors included urinary incontinence, tremors, abnormal stretching gait, hunched posture, back arching, and repeated licking of the thoracoabdominal region. To eliminate interference from the experimenter’s cage handling and blood collection procedures on the remaining rats, observations were performed once daily (9:20-9:50 am), 20 minutes after JV blood collection. The observer entered the housing room gently and allowed the rats to acclimate to their presence for 5 minutes (to avoid stress induced by sudden appearance). Subsequently, the behaviors of rats in each group were continuously recorded for 5 minutes beside the cages, with real-time documentation of the frequency of normal behaviors and pain/stress-related behaviors using a predesigned behavioral scoring sheet (including behavioral types and occurrence frequencies).

Histopathological analysis.

To assess vascular and tissue injury caused by jugular sampling, histopathologic examinations of cervical skin and blood vessels were performed using H&E staining. At the end of the repeated sampling experiment, 4 rats from each group were randomly selected by a technician blinded to group assignment. Animals were first anesthetized with Zoletil 50 (30 mg/kg IP injection; Virbac, Carros, France). Anesthetic efficacy was verified by the absence of eyelid closure response to gentle corneal stimulation (corneal reflex) and lack of limb withdrawal in response to hindlimb pinch. A 1- to 2-cm midline abdominal incision was made to expose the abdominal aorta, followed by exsanguination for euthanasia to avoid blood contamination of tissue samples. Blood was collected into vacuum tubes (Bangbiao Medical Equipment, Shanghai, China), and fur over the left cervical and thoracic regions was shaved. A 5-mm × 5-mm area centered 1 cm above the clavicle over JV (Figure 1C-b) was excised using sterile scissors, ensuring that the jugular vessel and adjacent tissues were intact. Samples were fixed in 10% formalin (Kangnaixin Biomedical Technology, Zhongshan, China) for 72 hours, embedded, and sectioned transversely at 3 μm. Sections were stained with H&E and examined under a microscope (Nikon, Shinagawa, Japan) by a blinded pathologist. Quantitative analysis of hemorrhagic area in the skin and vascular-muscular tissue was performed using ImageJ software.

Serum biochemistry.

Serum biochemical analyses were performed to evaluate the influence of the sampling methods on liver, kidney, lipid, and glucose metabolism parameters, which are commonly assessed in pharmacological and toxicological studies.^19,20 After completion of the repeated sampling experiment, 0.3 mL of blood was collected from the tail vein (Control group) or JV (TM and NM groups). Samples were left at room temperature for 2 hours and centrifuged at 3,000 rpm for 15 minutes to obtain serum. Biochemical parameters were measured using a Cobas c311 analyzer (Roche, Basel, Switzerland), including AST, ALT, total bilirubin (TBil), direct bilirubin (DBil), total protein (TP), albumin (ALB), albumin/globulin ratio (A/G), globulin (GLB), triglycerides (TG), total cholesterol (TC), high-density lipoprotein cholesterol (HDL-C), low-density lipoprotein cholesterol (LDL-C), BUN, creatinine (Cr), uric acid (UA), glucose (GLU), sodium (Na⁺), potassium (K⁺), chloride (Cl⁻), creatine kinase (CK), and LDH. In addition, serum or plasma was collected after the first and last blood collections (day 1 and day 7) during repeated sampling. Hemolysis in each group was observed by photography, and hemoglobin (HGB) levels were quantified using a BC-5120 automated hematology analyzer (Mindray, Shenzhen, China).

Coagulation function assays.

Coagulation testing was performed to determine whether sampling methods affected hematologic stability, as such assays are widely applied in hematologic disease models and preoperative evaluations.^21,22 After repeated sampling, 0.2 mL of blood was collected from the tail vein (Control) or JV (TM and NM groups) into sodium citrate tubes (Bangbiao Medical Equipment, Shanghai, China). Plasma was separated by centrifugation at 3,000 rpm for 15 minutes, and coagulation parameters were analyzed using a CA-1500 coagulation analyzer (Sysmex, Kobe, Japan). The indices included prothrombin time (PT), prothrombin percentage activity, international normalized ratio of PT (PT-INR), activated partial thromboplastin time (APTT), fibrinogen (FIB) concentration, and FIB time.

Hematologic parameters.

CBC analyses were performed to further assess the impact of sampling on hematologic parameters, which are of diagnostic value in infection, anemia, inflammation, and hematologic disease models.²³ Following repeated sampling, 0.2 mL of blood was collected from the tail vein (Control) or JV (TM and NM groups) into EDTA-K2 anticoagulant tubes. Samples were analyzed within 10 minutes using a BC-5120 automated hematology analyzer. Parameters included WBC, neutrophils (Neu), lymphocytes (Lym), neutrophil percentage, lymphocyte percentage, RBC, HGB, Hct, mean corpuscular hemoglobin concentration (MCHC), platelets (PLT), MPV, platelet distribution width (PDW), plateletcrit (PCT), platelet large cell count (P-LCC), and platelet large cell ratio (P-LCR).

Intraperitoneal glucose tolerance test.

To further evaluate the applicability of JV sampling in pharmacokinetic and toxicokinetic studies, an intraperitoneal glucose tolerance test (IPGTT) was performed, using glucose as a representative metabolic substrate to compare blood glucose profiles obtained with different sampling methods. After completion of the repeated sampling experiment, the remaining rats were allowed a 2-week recovery period under acclimatization. Animals were then weighed and randomly allocated into 3 groups (n = 4 per group): Control, TM, and NM groups. Rats were fasted for 12 hours (8:00 pm-8:00 am) before the experiment but had free access to water. A 20% glucose solution was prepared in PBS and administered intraperitoneally at 5 mL/kg (equivalent to 2 g/kg glucose). Blood samples (0.05 mL) were collected at 0, 15, 30, 60, 120, and 240 minutes postinjection according to group-specific sampling methods: tail tip vein for the Control group and JV for the TM and NM groups. Hemostasis was achieved by compression with a sterile cotton ball for 3-5 seconds after each puncture. Blood glucose concentrations were measured using a portable glucometer and test strips (Roche, Basel, Switzerland). Following the experiment, all animals were returned to normal housing conditions.

Acute kidney injury model validation.

To validate the applicability of the sampling methods in disease models, a cisplatin-induced acute kidney injury (AKI) rat model was established, and the 2 JV techniques were compared in terms of data stability and consistency. After recovery from the IPGTT, the remaining rats were acclimatized for 1 week, weighed, and randomly divided into 3 groups (n = 4 per group) using the Excel RAND () function: Control, TM, and NM groups. One day before cisplatin administration, 0.2 mL of blood was collected from the tail vein of all animals, allowed to clot at room temperature for 2 hours, and centrifuged at 3,000 rpm for 15 minutes. Serum Cr and urea concentrations were measured using a Cobas c311 analyzer (Roche, Basel, Switzerland) to establish baseline values. Cisplatin was prepared in physiologic saline containing 1% DMSO at a dose of 15 mg/kg (5 mL/kg) according to a published protocol²⁴ and administered intraperitoneally to induce AKI. After 48 hours, blood samples were collected from the tail vein (Control) or JV (TM and NM groups), and serum Cr and urea concentrations were measured to assess renal injury.

To directly compare the effects of sampling methods on biochemical outcomes, additional samples were collected at 48 hours from the Control group using both the traditional (TMJV-CIS-48h) and new (NMJV-CIS-48h) methods, followed by comparative analysis of Cr and urea levels. After completion of all sampling procedures, rats were euthanized using a CO₂ chamber (flow range: 0-25 L/min; Siargo, Chengdu, China). The CO₂ displacement rate for euthanasia was maintained in a range of 30%-70% container volume per minute, which ensured a humane and painless death in compliance with animal welfare standards.²⁵

Statistical analysis.

Data were analyzed using SPSS 25.0 (SPSS). Continuous variables are expressed as mean ± SD. Before parametric testing, normality was assessed with the Shapiro-Wilk test and homogeneity of variance with the Levene test. When assumptions were met, independent sample t tests (2 groups) or one-way ANOVA (multiple groups) were used, followed by least significant difference (equal variances) or Dunnett T3 (unequal variances) post hoc comparisons as appropriate. When assumptions were violated, Mann-Whitney U tests were applied. For categorical data, results were analyzed using χ² tests. Statistical significance was set at P < 0.05.

Results

Superior convenience of the new technique.

In accordance with our exclusion criteria, 24 rats were included in the present study. During the 7-day, 13-time-point protocol, body weight remained stable across all groups without significant differences between groups (Figure 2B). Hemolysis was reduced in the NM group compared with the TM group, as reflected by clearer serum/plasma supernatants (Figure 2C) and lower HMG levels (Figure 2D). Single sampling time was significantly shorter in the NM group at most time points (Figure 2E), and the overall mean sampling time was approximately halved compared with the TM group (34.7 ± 19.1 seconds compared with 71.1 ± 36.0 seconds; P < 0.001; Figure 2F). Likewise, puncture attempts were significantly fewer in the NM group (1.4 ± 0.6 compared with 2.6 ± 1.1; P < 0.001; Figure 2G and H), and the success rate reached 100% at all time points, compared with 69.2% ± 13.1% in the TM group (Figure 2I and J).

Reduced pain- and stress-related behaviors in the new technique group.

To assess the impact on animal welfare, behavioral responses were systematically observed during the repeated sampling protocol. Normal behavior frequencies did not differ significantly between groups (Figure 3A), with rearing being most common, followed by grooming, and, less frequently, feeding and drinking. In contrast, pain- and stress-related behaviors increased progressively in the TM group, peaking at day 7, and were significantly more frequent than in the NM group (P < 0.05) (Figure 3B and C). Licking the chest and abdomen and arching the back occurred most frequently, followed by hunched posture and staggering (data not shown). Cumulative analysis confirmed a 10.72% reduction in pain-related behaviors in the NM group compared with the TM group (Figure 3D).

Figure 3.

Behavioral Outcomes Associated with JV Sampling. (A) Frequency of normal behaviors. (B) Frequency of pain-related behaviors. (C) Representative images of normal and pain-related behaviors: (a) drinking; (b) grooming; (c) low and high posture rearing; (d) feeding; (e) thoracoabdominal licking; (f) back arching; (g) hunched posture; and (h) abnormal gait. (D) Cumulative behavioral counts. Data are presented as mean ± SD (n = 8); *P < 0.05 vs. TM group (pain-related behaviors); independent-samples t test.

Reduced hemorrhagic injury at the puncture site.

Histopathological analysis was performed to evaluate tissue damage at the sampling sites. Control animals displayed intact vascular and skin structures (Figure 4A and C). In contrast, the TM group exhibited extensive vascular hemorrhage, adjacent muscle disruption, and subcutaneous bleeding, with occasional inflammatory infiltration. The NM group showed largely preserved vessel morphology and minimal bleeding. Quantitative analysis confirmed that hemorrhagic areas in both vascular (2,293,889.0 ± 923,276.5 μm² compared with 8,642,455.3 ± 4,584,711.9 μm²; P < 0.05) and skin tissues (69,306.0 ± 43,345.3 μm² compared with 1,799,133.5 ± 1,282,991.3 μm²; P < 0.05) were significantly smaller in the NM group compared with the TM group (Figure 4B and D).

Figure 4.

Histopathological Analysis of Cervical Vessels and Skin Following JV Sampling. (A) Representative H&E-stained sections of cervical vessels and muscle tissue; magnifications 40× and 160×. Lower panels show enlarged regions indicated by black boxes in the upper panels. Red dashed lines and arrows indicate the JV and JV bleeding (JVB). (B) Quantitative analysis of hemorrhagic area in vascular and muscle tissue. (C) Representative H&E-stained sections of cervical skin; magnifications 40× and 160×. Lower panels show enlarged regions indicated by black boxes. Red dashed lines and arrows indicate skin tissue bleeding (STB). (D) Quantitative analysis of hemorrhagic area in skin tissue. Data are presented as mean ± SD (n = 4); ^##P < 0.01 vs. Control group; *P < 0.05 vs. TM group; one-way ANOVA or Mann-Whitney U test.

Greater stability of serum biochemical parameters with the new technique.

Serum biochemical analysis showed that the NM group exhibited stable parameters comparable to those of the Control group (Figure 5A-P), whereas the TM group displayed significant alterations. In particular, CK and LDH were markedly elevated in the TM group compared with the Control group (P < 0.01). By contrast, CK and LDH levels in the NM group (291.33 ± 199.79 U/L and 127.8 ± 63.4 U/L, respectively) were substantially lower than in the TM group (878.83 ± 818.04 U/L and 229.0 ± 86.0 U/L, respectively), approaching normal ranges (Figure 5Q and R). Although minor variations were observed among groups, all parameters remained within or close to the physiologic reference ranges for SD rats.

Figure 5.

Effects of JV Sampling on Serum Biochemical Indices. (A) AST, (B) ALT, (C) TBil, (D) DBil, (E) ALB, (F) A/G ratio, (G) TG, (H) TC, (I) HDL-C, (J) LDL-C, (K) BUN, (L) Cr, (M) GLU, (N) Na⁺, (O) K+, (P) Cl⁺, (Q) CK, and (R) LDH. Data are presented as mean ± SD (n = 6); ^##P < 0.01 vs. Control group; *P < 0.05 vs. TM group; one-way ANOVA or Mann-Whitney U test.

Preservation of coagulation function with the new technique.

Coagulation assays revealed that the NM group maintained stable coagulation indices comparable to the Control group, whereas significant differences were observed for the TM group (Figure 6). Specifically, the TM group exhibited shortened PT time (8.8 ± 0.3 seconds compared with 9.5 ± 0.3 seconds in Control; P < 0.05) and decreased PT-INR (P < 0.05), while FIB concentration (2.5 ± 0.8 g/L) and FIB time (7.0 ± 1.4 seconds) were significantly altered relative to the Control group. In contrast, the NM group showed values closer to baseline, with significantly reduced deviations in FIB concentration (1.6 ± 0.1 g/L) and FIB time (9.6 ± 0.5 seconds) compared with the TM group (P < 0.05). Despite minor intergroup differences, all values remained within normal physiologic ranges.

Figure 6.

Effects of JV Sampling on Coagulation Parameters. (A) PT time, (B) PT activity (%), (C) PT-INR, (D) APTT, (E) fibrinogen (FIB) concentration, and (F) fibrinogen (FIB) time. Data are presented as mean ± SD (n = 4); ^#P < 0.05 vs. Control group; *P < 0.05 vs. TM group; one-way ANOVA or Mann-Whitney U test.

Stability of hematologic parameters with the new technique.

CBC analysis (Table 1) demonstrated that the NM group maintained hematologic parameters more consistent with the Control group, while the TM group exhibited significant alterations. RBC-related indices (RBC, HGB, and Hct) were significantly reduced in the TM group compared with Control group (P < 0.01), but values remained comparable to the Control group in the NM group, with values significantly higher than those observed in the TM group (P < 0.01). WBCs and their subsets (WBC, Neu, and Lym) did not differ significantly among groups. For platelet indices, PDW was significantly elevated in the TM group relative to the Control group (P < 0.05), whereas PDW in the NM group did not differ from Control. Overall, all hematologic indices remained within or close to the established reference ranges for SD rats, and the NM group demonstrated superior stability compared with the TM group.

Table 1.

Hematologic Parameters in Rats Across Groups

Hematologic parameters	Control	TM	NM
WBC, ×103/μL	7.70 ± 1.51	8.17 ± 1.95	7.37 ± 1.12
Neu, ×103/μL	1.18 ± 0.21	1.56 ± 0.92	1.11 ± 0.29
Lym, ×103/μL	6.50 ± 1.42	6.56 ± 1.22	6.24 ± 1.04
Neu, %	15.53 ± 3.01	18.19 ± 6.44	15.01 ± 3.65
Lym, %	84.10 ± 3.06	81.23 ± 6.65	84.61 ± 3.65
RBC, ×106/μL	6.21 ± 0.20	5.31 ± 0.42##	5.92 ± 0.33**
HGB, g/L	131.75 ± 4.50	117.38 ± 10.17###	130.50 ± 4.78**
Hct, %	37.54 ± 1.45	33.75 ± 2.95##	37.25 ± 1.50**
MCHC, g/L	351.13 ± 3.00	347.75 ± 4.74	350.00 ± 2.83
PLT, ×103/μL	899.38 ± 150.50	940.38 ± 334.92	1,006.00 ± 176.97
MPV, fL	6.59 ± 0.36	7.26 ± 0.99	6.51 ± 0.20
PDW, %	14.94 ± 0.09	15.13 ± 0.23#	14.94 ± 0.14*
PCT, %	0.59 ± 0.12	0.66 ± 0.20	0.65 ± 0.11
P-LCR, %	6.79 ± 1.69	10.61 ± 5.98	6.28 ± 0.91

Data are presented as mean ± SD (n = 8). Statistical analysis was conducted using one-way ANOVA, and nonparametric data were analyzed using Mann-Whitney U test.

^#P < 0.05, ^##P < 0.01, ^###P < 0.001 vs. Control; *P < 0.05, **P < 0.01 vs. TM.

Consistent application of the new technique in IPGTT and AKI models.

The applicability of the new technique was further validated in pharmacokinetic and disease model settings. In the IPGTT experiment, glucose concentrations in the NM group closely paralleled those of the Control group at all time points (0-240 minutes), with no significant differences in blood glucose levels or AUC values (2,083.3 ± 213.9 compared with 2,298.8 ± 139.0 mmol/L·min; P > 0.05) (Figure 7A and B). By contrast, the TM group has high variance at several time points and a significantly reduced AUC compared with Control (1,963.3 ± 170.1 mmol/L·min; P < 0.05; Figure 7C). In the AKI model induced by cisplatin (Figure 7D-G), BUN and Cr levels increased significantly across all groups (P < 0.01 or P < 0.001), confirming successful model establishment. No significant differences were observed among groups, and further comparison within the Control group using different sampling methods revealed consistent BUN and Cr results.

Figure 7.

Validation of JV Sampling in Pharmacokinetic and Disease Models. (A) Blood glucose concentrations at different time points in the IPGTT (n = 4). (B) Individual glucose concentration curves for each rat (n = 4). (C) AUC of glucose concentration–time profiles (n = 4). (D) Serum BUN levels before and after cisplatin-induced AKI (n = 3). (E) Comparison of BUN levels in the Control group using different sampling methods at 48 h postcisplatin (CV-CIS-48h, TM-CIS-48h, and IM-CIS-48h) (n = 3). (F) Serum Cr levels before and after cisplatin-induced AKI (n = 3). (G) Comparison of Cr levels in the Control group using different sampling methods at 48 h postcisplatin (n = 3). Data are presented as mean ± SD (n = 3-4); ^#P < 0.05, ^##P < 0.01, ^###P < 0.001 vs. Control group; one-way ANOVA or independent-samples t test.

Discussion

In this study, we describe a new technique that integrates anatomic features of the rat cervical vasculature with optimized restraint and vascular puncture procedures, enabling both single and repeated blood collection in conscious animals. Compared with the traditional approach, the new method demonstrated clear advantages in efficiency, safety, and blood sample stability. Collectively, these findings support the new technique as a valuable refinement over existing methods, with direct relevance for preclinical pharmacological and toxicological studies.

Conventional JV sampling often relies on subclavian puncture,^13–15 which provides large blood volumes but carries substantial risks due to the close anatomic proximity of subclavian vessels to the brachial plexus, anterior vena cava, thoracic cavity, and heart.¹⁴ Such approaches can lead to severe complications including hemorrhage, pneumothorax, and nerve injury, particularly in long-term sampling studies. Our histopathology and behavioral evaluations further confirmed these safety risks. To mitigate them, the puncture site in the new technique was shifted to the lower cervical vein, located above the clavicle and away from dense vascular and neural structures, thereby reducing procedural risk from an anatomic perspective. In addition, the new method employs coordinated use of the palm and fingers to stabilize the animal, providing a firm and reproducible position for vascular puncture in contrast to conventional scruff restraint. The puncture landmark, the midpoint between the most prominent point of the shoulder joint and the thoracic midline, is intuitive and readily identifiable and, when combined with standardized insertion angle and depth, ensures reliable venous access. These refinements explain the enhanced efficiency and reproducibility of the new technique. Importantly, maintaining animal comfort is critical to procedural success: gentle handling, stroking of the dorsal skin, and minimizing stress are essential, whereas improper restraint may induce agitation, bleeding, or procedural failure.²⁶ Adequate training and practice are therefore indispensable prerequisites for successful implementation.

Body weight, a fundamental indicator of animal health,²⁷ remained stable across all groups during repeated sampling, suggesting that the new method does not interfere with normal physiology. Hemolysis, a critical determinant of sample quality,²⁸ was substantially reduced in the new group, yielding clearer serum/plasma and lower free hemoglobin levels. Importantly, by comparing different methods of JV blood collection in rats, we demonstrated the advantages of our method in terms of convenience, safety, and sample quality stability (Table S1). Convenience, a decisive factor for widespread adoption, was reflected in markedly shorter sampling times, fewer puncture attempts, and higher success rates compared with traditional approaches. Published reports indicate success rates of approximately 80% for JV catheterization,¹² approximately 90% for subclavian sampling under anesthesia with a minimum time of 65 seconds,¹³ and approximately 95% for conscious subclavian sampling with mean times exceeding 2 minutes.¹⁵ By contrast, our new technique achieved a 100% success rate with a mean sampling time of 34.7 seconds, approximately half the time required by traditional approaches. These advantages are likely attributable to the optimized restraint strategy, precise puncture localization, and procedural standardization. Nevertheless, as the present results were obtained under specific conditions and experimental cycles, further validation in diverse experimental contexts is warranted.

Safety, which is closely linked to animal welfare, is a primary concern in repeated blood collection. Severe hemorrhagic injury can compromise study continuity and animal survival. Our behavioral and histopathologic assessments confirmed that the new technique markedly reduced pain behaviors and minimized tissue injury relative to the traditional approach. Sample stability is another essential prerequisite for reliable pharmacological and toxicological studies.^29–31 The new technique produced more stable serum biochemical, coagulation, and hematologic indices than the traditional method. Particularly noteworthy were CK and LDH, classic markers of tissue damage and stress,^32,33 which were significantly elevated in the TM group but not in the NM group. Similarly, high variance in PT, FIB, RBC, and HGB observed in the TM group was absent in the NM group. In terms of data correlation, histopathologic analysis showed more hemorrhagic injuries in the TM group. Increased blood loss after repeated sampling may have contributed to hematologic parameter differences, with all results mutually validating. This is likely due to greater operational stress and tissue damage associated with the TM technique. Together, these findings indicate that the new technique effectively mitigates stress responses while ensuring sample quality.

The applicability of the new technique was further confirmed in models simulating typical applications in pharmacology and toxicology.^10,34 In the IPGTT, blood glucose profiles obtained by the new technique closely matched those of tail vein sampling, whereas the traditional method significantly reduced AUC values. Similarly, in the cisplatin-induced AKI model, both methods produced consistent BUN and Cr values comparable to tail vein sampling, indicating reliable applicability to disease model studies.²⁴ Together, these results demonstrate that the new technique is suitable for frequent microsampling and disease model analysis, with performance comparable to or better than established approaches.

To our knowledge, this study represents the first systematic comparative evaluation of JV sampling techniques in rats, clarifying the advantages of the new approach. Nonetheless, limitations remain. First, corticosterone measurements were not performed to quantitatively assess stress responses. Second, no direct comparison was made with other commonly used methods, such as retro-orbital or submandibular sampling. In addition, the present work was limited to male SD rats, and applicability to female rats or other strains remains to be established. Future studies should address these aspects and evaluate long-term applications to further refine the method.

In summary, we have developed a new single-operator JV sampling technique for conscious rats, providing a convenient, safe, and stable method for obtaining high-quality blood samples. This technique offers reliable support for both short- and long-term pharmacological, toxicological, and pharmacokinetic studies, while simultaneously promoting animal welfare and contributing to refinement in line with the Replacement, Reduction, and Refinement (3Rs) principles.

Conclusion.

We developed a new JV sampling technique for conscious rats, integrating standardized precise puncture localization and bilateral alternating sampling. It enables a single operator to efficiently collect high-quality repeated blood samples without auxiliaries. Convenient, safe, and blood quality stable, it accelerates reliable sample and accurate data acquisition in short and long-term experiments.

Supplementary Materials

Table S1. ^{(24KB, docx)} Comparison of different rat jugular vein blood collection methods in terms of convenience, safety, and sample quality stability.

Conflict of Interest

The authors have no conflicts of interest to declare.

Funding

This work was supported by the Guangdong Provincial Basic and Applied Research Foundation (Grant No. 2022A1515140082).

Author Contributions

Mingqiang Wang: study design, experimentation, data analysis, and manuscript drafting. Juncheng Deng: resources, methodology, supervision, and manuscript revision. Youan Xue, Yingyong Wen, and Yongjiao Xie: experimentation and resources. Yi Jin: methodology and supervision. Jing Li: project administration, review, and editing.

Data availability

The original data supporting the findings of this study are available from the corresponding author upon reasonable request.