All procedures involving animals in this study were conducted in full compliance with national and institutional guidelines governing the ethical use of animals in research. The study protocol, including animal selection, handling, blood collection techniques, and sample processing, was reviewed and approved by the Institutional Animal Care and Use Committee of the Faculty of Veterinary Medicine, Chiang Mai University, Thailand (Approval No. FVM-ACUC; S8/2561). This approval confirms that the study adhered to the ethical principles outlined in the Animal Research: Reporting of In Vivo Experiments 2.0 guidelines and followed standards designed to minimize discomfort, stress, and risk to participating animals.

All elephants enrolled in the study were privately owned and housed in elephant camps that routinely collaborate with Chiang Mai University for health monitoring and welfare management. Before sampling, each elephant underwent a physical examination by a licensed veterinarian to confirm eligibility and ensure no underlying health conditions or infectious diseases were present. Blood samples were collected by trained veterinarians or experienced elephant handlers using minimally invasive techniques. Positive reinforcement (food rewards) was used to reduce stress during handling, and no sedation or coercive methods were used. The entire sampling process was completed within a short time frame to minimize disturbance.

The study did not involve experimental treatments, drug administration, or procedures likely to cause pain or long-term physiological effects. Only the volume of blood permitted under institutional welfare guidelines for large mammals was collected, and all animals returned immediately to their normal husbandry routines after sampling. No elephant experienced adverse effects during the study.

This ethical approval ensures that the work was conducted with the utmost regard for animal welfare, scientific integrity, and regulatory compliance, recognizing the cultural, conservation, and ecological importance of Asian elephants.

A total of 20 healthy Asian elephants, 10 males and 10 females, were selected from two elephant camps in Chiang Mai, Thailand. Elephants showing signs of illness or infection were excluded from the study. The animals ranged in age from 10 to 60 years. All elephants were fed a diet primarily of corn stalks and Napier grass (Pennisetum purpureum) and had unrestricted access to fresh water. They regularly participated in tourist-related activities, including saddle riding, bathing, and feeding. Physical examinations conducted by camp veterinarians confirmed their good health at the time of sampling.

Blood samples were collected once from an ear vein by trained elephant camp staff or veterinarians from Chiang Mai University in April 2018. Using a 1-inch, 18-gauge IV catheter and a 50-mL syringe, 30 mL of blood was collected between 10:00 a.m. and 12:00 p.m. Each collection was completed within 5 min, with elephants calmly restrained using positive reinforcement (food rewards). Fourteen milliliters of blood were transferred into 50-mL conical tubes preloaded with citrate phosphate dextrose (CPD) anticoagulant (C7165, Sigma-Aldrich, MO, USA) at a 1:7 ratio (2 mL CPD:14 mL blood). The remaining 16 mL were placed into red-top tubes (BD Vacutainer® Serum, USA) without anticoagulant and left undisturbed for 2 h to allow serum separation. All tubes were clearly labeled, kept on ice at approximately 4°C in a Styrofoam container, and transported immediately for processing within 4 h of collection.

Upon arrival at the laboratory, anticoagulated blood was centrifuged at 700 × g for 10 min at 4°C (Eppendorf 5810R, Germany) to obtain plasma, yielding 55%–60% of total blood volume. Plasma was aliquoted into four microcentrifuge tubes (2 mL each). Serum tubes were centrifuged at 252 × g for 10 min, yielding 40%–45% serum, which was aliquoted into eight microcentrifuge tubes (1 mL each).One plasma and two serum aliquots were analyzed immediately, while the remaining samples were stored at −20°C for analysis at 4, 8, and 12 months with temperature monitoring. Prior to testing, frozen plasma and serum were thawed in a 37°C water bath and analyzed within 2 h. Samples showing hemolysis or lipemia were excluded.

Plasma was used for fibrinogen and factor VIII assays because it contains intact clotting factors, which are preserved by anticoagulants. Serum, which lacks clotting factors due to coagulation during processing, was used for IgG and albumin assays to minimize assay interference.

Coagulation and protein concentrations were reported as mean ± standard deviation. A repeated-measures generalized linear model (GLM) was used to assess the effects of storage duration (0, 4, 8, and 12 months) on plasma protein stability. Model diagnostics included visual evaluation of Q-Q plots and residuals versus fitted values to confirm normality and homoscedasticity. Shapiro–Wilk tests were also performed. The GLM model was specified as: value ~ sex * month + (1 | elephantID). Post hoc pairwise comparisons were conducted using the multcompView package with Tukey-adjusted P values. Statistical significance was set at α = 0.05. All analyses were performed using R software (version 4.1.2; R Core Team, Vienna, Austria, 2023) [34].

The average concentrations of fibrinogen, factor VIII, IgG, and albumin in pre- and post-storage plasma samples stored at −20°C for 4, 8, and 12 months are summarized in Table 1. Statistical analyses showed that the month of collection was a significant predictor in all models, whereas sex did not significantly affect any coagulation or protein parameters (Table S1). Across storage durations, mean fibrinogen (F_3,54 = 11.46, p < 0.001), factor VIII (F_3,54 = 8.53, p < 0.001), IgG (F_3,54 = 39.30, p < 0.001), and albumin (F_3,54 = 26.87, p < 0.001) concentrations differed significantly.

Fibrinogen concentrations remained stable across storage periods. No significant differences were observed between fresh plasma and plasma stored for 8 months (248.63 ± 26.37 mg/dL, p = 0.165) or 12 months (248.69 ± 26.35 mg/dL, p = 0.172). These results indicate that fibrinogen remains stable when stored at −20°C for up to 12 months.

Factor VIII activity declined markedly over time. Significant reductions were observed in samples stored for 8 months (750.30 ± 170.72 IU/dL, p < 0.001) and 12 months (759.39 ± 162.24 IU/dL, p < 0.001) compared with fresh plasma. Overall, factor VIII activity decreased by approximately 16% after 12 months of storage, though values remained within clinically acceptable limits.

Mean IgG concentrations increased substantially in frozen plasma samples across all storage durations. Significant differences were observed between fresh and stored plasma at 4 months (5.79 ± 0.65 g/dL, p = 0.003) and 8 months (6.81 ± 1.09 g/dL, p < 0.001), with a continued upward trend at 12 months (6.90 ± 1.14 g/dL, p < 0.001). By 12 months, IgG levels had increased by approximately 37%.

Albumin levels also increased significantly during storage. Compared with fresh plasma, albumin concentrations were higher at 8 months (3.58 ± 0.33 g/dL, p = 0.029) and at 12 months (4.02 ± 0.39 g/dL, p < 0.001). Overall, albumin levels rose by approximately 21% after 12 months of frozen storage.

The temporal trends in fibrinogen, factor VIII, IgG, and albumin concentrations across all storage intervals (0, 4, 8, and 12 months) are shown in Figure 1, illustrating the stability or variation of each parameter during −20°C storage.

Unlike studies in canine and human medicine, this investigation represents the first controlled assessment of long-term protein stability in FFP derived from Asian elephants. These findings provide species-specific insights essential for developing emergency therapeutic strategies for EEHV-HD. Establishing evidence-based plasma storage guidelines is critical for creating the first plasma banking standards for captive elephants, enabling rapid intervention during hemorrhagic crises. Plasma transfusion plays a vital therapeutic role by supplying hemostatic proteins, mitigating hypovolemic shock, supporting immune function, and reducing the risks of red blood cell sensitization and volume overload. Although human FFP can be stored for up to 12 months at −20°C under American Association of Blood Banks guidelines, these standards may not directly apply to elephants because of species-specific differences in plasma protein stability.

This study demonstrated a significant increase in fibrinogen concentrations after 4 months of frozen storage at −20°C, with non-significant decreases at 8 and 12 months compared with fresh plasma. These findings align with canine studies reporting stable fibrinogen levels after 30 days at −30°C [35] and with human studies showing no significant differences in fibrinogen when stored at −20°C or −70°C for 4 months [27]. Storage at ultra-low temperatures (−196°C) may provide even better preservation [27]. The Clauss assay used here is widely implemented in clinical settings [33], although discrepancies between Clauss and fibrinogen-derived methods have been documented [36, 37]. While freezing and storage can strongly influence PT and activated partial thromboplastin time (aPTT), fibrinogen appears less affected, with smaller changes reported at −70°C than at −20°C [27].

Factor VIII concentrations decreased progressively during storage at −20°C, consistent with studies of canine plasma stored at −30°C for up to 12 months [26] and observations of declining factor VIII activity in canine plasma stored at −80°C [38]. As a labile protein, factor VIII is susceptible to deterioration during freezing, thawing, and long-term storage. The automated assay used may also contribute to measurement variability. Thawing at 37°C followed standard protocols [39], and prior studies show no detrimental effect on fibrinogen or factor VIII when thawed at 37–45°C [40] or across a wider temperature range (22°C–60°C) [41]. Despite the decline, factor VIII activity remained within clinically acceptable limits, supporting the potential clinical use of stored elephant FFP.

Storage at −20°C increased concentrations of IgG and albumin, likely due to cryoconcentration or subtle fluid loss during freezing. These elevated protein levels may prove beneficial in mitigating the hypoproteinemia characteristic of EEHV-HD. The successful adaptation of human-grade laboratory assays (Clauss assay, one-stage clotting assay, and colorimetric tests) to elephant plasma demonstrates methodological innovation and supports cross-species assay applicability.

The selected storage durations (4, 8, and 12 months) were informed by prior canine FFP studies showing retained therapeutic activity after 3–12 months of storage [26]. Elephant plasma was aliquoted into small volumes (1–2 mL) to promote rapid freezing, in contrast to clinical plasma bags (200–250 mL), which freeze more slowly and may compromise factor stability. This study’s longitudinal design and use of repeated-measures GLM provide a rigorous statistical evaluation rarely achieved in wildlife transfusion research. Elephants may possess unique plasma protein stability characteristics, reflecting evolutionary adaptations in large mammals. Although comparative data are limited, the biological plausibility adds originality and relevance to the study.

This study is limited by its modest sample size and assessment of a single storage temperature (−20°C). Future research should examine lyophilized [32, 42] and cryopreserved plasma at −80°C and −196°C, extend storage durations, and use larger aliquot volumes to better replicate clinical banking conditions. Additional clotting factors (e.g., II, VII, IX, X, and vWf) and coagulation parameters (PT, aPTT) should be evaluated to expand understanding of plasma stability. Clinical outcome studies involving EEHV-HD calves transfused with stored FFP are essential to validate therapeutic effectiveness.

Fibrinogen concentrations remained stable over 12 months of storage at −20°C, while factor VIII declined by 16% and IgG and albumin increased by 37% and 21%, respectively. Despite these changes, all parameters remained within clinically acceptable ranges for potential therapeutic use. Further investigations across varied storage conditions and longer durations are needed to refine species-specific guidelines. This study contributes to One Health efforts [43] by integrating wildlife medicine, transfusion science, and conservation biology to establish foundational protocols for elephant blood banking. The findings support the development of emergency FFP reserves for endangered megafauna, including rhinoceroses and tapirs, advancing conservation physiology and improving preparedness for critical care interventions.