Morphological and quantitative analysis of leukocytes in free-living Australian black flying foxes (Pteropus alecto).

The black flying fox (Pteropus alecto) is a natural reservoir for Hendra virus, a paramyxovirus that causes fatal infections in humans and horses in Australia. Increased excretion of Hendra virus by flying foxes has been hypothesized to be associated with physiological or energetic stress in the reservoir hosts. The objective of this study was to explore the leukocyte profiles of wild-caught P. alecto, with a focus on describing the morphology of each cell type to facilitate identification for clinical purposes and future virus spillover research. To this end, we have created an atlas of images displaying the commonly observed morphological variations across each cell type. We provide quantitative and morphological information regarding the leukocyte profiles in bats captured at two roost sites located in Redcliffe and Toowoomba, Queensland, Australia, over the course of two years. We examined the morphology of leukocytes, platelets, and erythrocytes of P. alecto using cytochemical staining and characterization of blood films through light microscopy. Leukocyte profiles were broadly consistent with previous studies of P. alecto and other Pteropus species. A small proportion of individual samples presented evidence of hemoparasitic infection or leukocyte morphological traits that are relevant for future research on bat health, including unique large granular lymphocytes. Considering hematology is done by visual inspection of blood smears, examples of the varied cell morphologies are included as a visual guide. To the best of our knowledge, this study provides the first qualitative assessment of P. alecto leukocytes, as well as the first set of published hematology reference images for this species.

Introduction

Several bat-borne viruses are of human health concern when cross-species transmission events, or spillovers, occur. Presently, the majority of emerging human pathogens have an origin in animals, with many arising from wild mammalian species [1]. For a pathogen to spill over from its animal host into a human, multiple factors must align in space and time. This encompasses both ecological and epidemiological conditions, including changes in the health of the host that benefit pathogen shedding [2]. Thus, a reservoir host must shed the pathogen for it to potentially encounter a susceptible human to infect. As a result, spillover events are closely linked to the ability of a reservoir host’s immune system to control infection, replication, and eventually pathogen shedding. In the case of bats, increased shedding of several viral species that infect them have been associated with periods of energetic or physiological stress [3-6]. The characterization of “health” is difficult in free-ranging wildlife, largely due to a lack of suitable metrics in wild animals with an unknown medical history. In this context, hematological analyses are frequently used as one of a suite of metrics to assess health, especially because of the role of white blood cells in the response of organisms to infections and other physiological stressors, which translates into changes in the numbers and proportions of different cell types [7]. If hematological changes have predictive power regarding infection outcomes, gaining a better understanding of bat hematology may help us to understand viral shedding, and therefore predict spillover events [8].

Pteropus spp. bats (colloquially known as flying foxes) are an important focus of bat-borne pathogen research [9]. Bat species within this genus are known or suspected reservoir hosts of several zoonoses, including Hendra virus, Nipah virus, Australian bat lyssavirus, and Menangle virus [6, 10–14]. In recent years in eastern Australia, flying foxes have increasingly occupied roosts within urban and peri-urban agricultural areas that offer predictable but low quality food resources [4]. These behavioral changes may facilitate contacts between bats and humans, and bats may also be experiencing poor health due to low food quality. As contact between flying fox populations, domestic animals, and humans increases, so does the risk of transmission of zoonotic infectious pathogens [3, 4, 6, 7]. One species of interest is Pteropus alecto (black flying fox) which play important ecological roles as pollinators and seed dispersers, and are endemic to Australia, Indonesia, and Papua New Guinea [15]. Despite the role of P. alecto as reservoirs of Hendra virus, relatively few studies have tried to characterize the physiological condition of clinically “healthy” individuals to serve as a baseline for health assessments. In the case of hematology, studies are generally conducted using automated cell counters. This approach, although highly efficient in terms of human hours and samples processed, can be impractical due to the logistics of using cytometers in remote field conditions, and the higher sample volume requirements for automated counting relative to slide preparation. Automated cell counters can also present errors associated with variability in cell types among species. In this context, manual assessment of cell morphology is required to establish a baseline of leukocyte characteristics. In this study, we aimed to address knowledge gaps regarding detailed morphological characterizations of the leukocytes of “healthy” black flying foxes. Here we provide an in-depth, microscopy-based characterization of P. alecto leukocytes, including morphological descriptions, reference photographs, and quantification of proportions of cell populations. The atlas of images provided in this study is aimed at facilitating the identification of P. alecto leukocytes in future studies and clinical settings.

Materials and methods

Animal ethics permits

This study was carried out in accordance with guidelines for animal care and handling under Griffith University Animal Ethics Committee (Approval ENV/10/16/AEC) and Montana State University IACUC Committee (#201750).

Animal capture and sample collection

P. alecto were captured between June 2018 and July 2020 at two roost sites in Queensland, Australia: one in Redcliffe and one in Toowoomba (S1 Table). Redcliffe is a coastal suburb approximately 30 kilometers northeast of Brisbane. Toowoomba is a small inland city 700 m above sea level in the Great Dividing Range, approximately 120 kilometers west of Brisbane. These roost locations are spaced approximately 160 kilometers apart and are continuously occupied by P. alecto [16]. Bats were captured, examined, sampled, and released within their roost site.

Bats were captured pre-dawn in mist nets and anesthetized by a veterinarian or veterinarian-trained technician using 5% isoflurane in oxygen at 800 mL/min, followed by administration of 1.5% isoflurane in oxygen at the same rate once animals were fully anesthetized. Physical examination was conducted to identify age, sex, and body condition, as well as any macroscopic injuries or abnormalities. Age was estimated as juvenile (less than 1 year old), subadult (pre-reproductive, ~1–2 years old), and adult (greater than ~2 years old) based on morphometric measurements of forearm length, body mass, tooth appearance, and reproductive maturity [12, 17]. Distinction between adults and subadults was made based on penis and testes size and development in males, and pregnancy (via abdominal palpation) or evidence of past suckling (based on nipple protrusion and balding around nipples) in females.

All bats were marked by painting the claws on one hind limb with colored nail lacquer to identify recently captured bats and avoid resampling during the consecutive days of capture sessions. Bats captured from May 2019 onwards were also marked with a subcutaneous RFID Passive Integrated Transponder (PIT tag, or ‘microchip’; ZD Tech Group, China) inserted between the scapulae. After sampling, each bat was monitored for recovery from anesthesia for at least 30 minutes; good grip ability and airway stability were confirmed before release.

Prior to PIT tag insertion, a maximum of 2.5 mL of blood was drawn from the cephalic or uropatagial vein, with samples not exceeding 0.6% of body mass. Blood smears were prepared on-site upon sample collection using blood drawn directly from the syringe, without the use of anticoagulant agents. Multiple smears were made for each bat when blood volume allowed. Smears were dried at ambient temperature and fixed in 100% methanol for three minutes. The samples were then stored at room temperature and out of UV light for up to two years until analysis at Montana State University (Import Permit No. 20200728-2504A).

Hematological analysis

Blood smears were stained with the commercial Romanowsky stain variant DipQuik (Jorgensen Laboratories, Loveland, CO, 80538, USA). Shape, length, texture, and cell monolayer of each smear were examined for quality, and only medium and high-quality smears were analyzed. Smears with uneven distribution of leukocytes, damage or poor stain quality, and high numbers of reactive, unidentifiable leukocytes were not analyzed.

The morphological characteristics of leukocytes, erythrocytes, and thrombocytes in the monolayer were assessed using an AmScope E5 Biological Series microscope. Standard leukocyte differentials were performed manually by counting 100 different leukocytes in the monolayer of each smear. Up to three differential counts (up to 300 leukocytes total) were performed on each smear to ensure at least 2/3 of the monolayer was examined. To avoid inconsistencies in identification of cells that could bias results, a single laboratorian conducted all smear examinations.

The diameters of 355 cells from a subset of 15 individuals sampled at both roost sites (n Redcliffe = 7; n Toowoomba = 8) were measured using an AmScope E5 series biological microscope and 5.1 MP USB3.0 Aptina Color CMOS Microscope Digital Camera. 134 erythrocytes, 80 neutrophils, 70 lymphocytes, 30 monocytes, and 28 eosinophils were measured, and cells which were extremely oblong, distorted, or abnormal in appearance were excluded. The camera software was calibrated at all magnification levels directly prior to measuring cells. The overall means, medians, and ranges for the diameter of each cell type were then calculated.

Reference images were taken at 600x or 1000x magnification on a Nikon Eclipse 80i microscope or an AmScope LED E5 Biological Series microscope equipped with a 5.1MP USB3.0 Aptina Color CMOS Microscope Digital Camera. Representative images of each cell type and repeatedly noted unique morphologies were taken from the highest quality smears to maximize image quality.

Quantitative analysis

All data manipulation and visualizations were performed in R version 1.4.1106 using the packages tidyverse and ggplot2 [18-20]. Considering the low number of individual bats from which good quality smears were obtained, we opted to summarize the data providing inter-quartile ranges per sex and using only adult individuals (n = 134/134). Individual samples from both roosts were grouped to increase sample sizes. When differentials were performed on more than one smear from the same individual bat, the differentials were averaged so that each bat accounted for only one datapoint in the final dataset. Histograms were used to assess the normality of the data, and natural log-transformations in the form ln(x+1) were applied to the monocyte and eosinophil counts to facilitate visualizations by adjusting for skewness. Given the descriptive focus of this paper and the lack of absolute numbers of cells from which to draw robust statistical comparisons, no statistical analyses were performed.

Results

The final dataset used for this study consisted of data collected from 134 adult P. alecto captured at Redcliffe and Toowoomba, Queensland, Australia at 16 different catching sessions between June 2018 and July 2020 (S1 Table). All bats used in our analysis appeared clinically healthy at the time of capture with no obvious indications of disease; however, minor injuries or abnormalities, including wing tears, abrasions, evidence of healed wounds, and mild dermatitis were observed on multiple individuals (n = 24; 13 females and 11 males). These individuals were excluded from calculations of the prevalence of morphological variations. A single individual was excluded from our analyses due to an exceptionally high neutrophil count. The anomaly was confirmed in two smears prepared from the individual and the reason for the elevated value remains unclear. An additional individual was also excluded from analysis due to pronounced emaciation.

Morphological descriptions

Neutrophils

Neutrophils were the dominant leukocyte type, comprising over 50% of leukocytes in 92.72% of adult bats (n = 102/110) (Tables 1 and 2 and Fig 1). Neutrophils (Fig 2A–2H) were characterized by pale purple, pink, or grey cytoplasms and dark purple lobulated nuclei. There was high variability in the number of lobes observed, with numerous hyposegmented cells (less than three lobes) and hypersegmented cells (greater than four lobes) observed (Fig 2G and 2H). Low numbers of activated neutrophils were observed throughout the sample set. Neutrophils were generally observed to be smaller in diameter than monocytes, but larger than lymphocytes (Table 3 and Fig 3). Neutrophils were primarily agranular with smooth cytoplasmic textural appearances, or hypogranular with slightly textured and darker staining cytoplasm. Granular neutrophils were observed in 3.63% of individuals (n = 4/110). In these cells, granules were large, round, purple to reddish pink in color, and dispersed throughout the cytoplasm at a low density (Fig 2C). Where present, granular neutrophils accounted for 1.4–4% of the total neutrophil population. Most granular neutrophils observed were banded or hyposegmented. Granular or agranular band neutrophils were observed in 34.54% of individuals (n = 38/110) (Fig 2A–2C).

Table 1

Leukocyte profiles of P. alecto with no observed injuries.

	IQR		Median		Range
Leukocyte Type	Female (n = 55)	Male (n = 55)	Female (n = 55)	Male (n = 55)	Female (n = 55)	Male (n = 55)
Neutrophils	55.0–74.8	55.0–71.5	70.0	65.0	26.0–91.0	37.0–88.0
Lymphocytes	19.0–36.8	22.5–33.5	25.0	30.0	9.0–74.0	8.5–51.0
Eosinophils	0.0–3.5	0.3–6.0	1.0	2.0	0.0–28.0	0.0–26.0
Monocytes	2.0–6.0	2.0–5.3	4.0	4.0	0.0–10.0	0.0–14.0

Interquartile ranges, median values, and overall ranges for each observed leukocytic cell type per 100 cells counted for Pteropus alecto on which no injuries were observed.

Table 2

Leukocyte profiles of P. alecto with observed mild/minor injuries.

	IQR		Median		Range
Leukocyte Type	Female (n = 13)	Male (n = 11)	Female (n = 13)	Male (n = 11)	Female (n = 13)	Male (n = 11)
Neutrophils	48.0–69.0	60.5–81.5	62.0	69.3	29.0–81.0	46.0–91.0
Lymphocytes	31.0–43.0	11.0–34.8	31.0	23.0	14.0–55.0	7.5–46
Eosinophils	0.0–2.0	0.0–1.0	1.0	0.3	0.0–12.5	0.0–5.5
Monocytes	3.0–7.0	2.8–7.0	5.0	5.0	0.5–9.0	0.0–9.0

Interquartile ranges, median values, and overall ranges for each observed leukocytic cell type per 100 cells counted for Pteropus alecto on which injuries were observed.

Fig 1

Box and whisker plots displaying the number of neutrophils (A), lymphocytes (B), eosinophils (C) and monocytes (D) observed per 100 cells counted in individuals on which injuries were noted (blue, n = 24) compared to individuals with no evident injuries (red, n = 110). Values of cell populations present in low abundances were log-transformed as ln(x+1) for visualization purposes.

Fig 2

Images taken at 600x or 1000x magnification depicting the variable morphologies of each leukocyte lineage as well as red blood cells and platelets.

(A, B) band neutrophils, (C) granular band neutrophil, (D-F) neutrophils, (G, H) hypersegmented neutrophils, (I-L) lymphocytes, (M) reactive lymphocyte, (N) binuclear lymphocyte, (O-P) large granular lymphocytes, (Q-R) band eosinophils, (S-X) eosinophils, (Y-Ff) mature monocytes, (Gg) erythrocytes, (Hh-Ii) platelet clumps, (Kk) echinocytes, (Ll) erythrocytes with low central pallor, (Mm) dacrocytes, (Nn) erythrocyte anisocytosis.

Table 3

Cell diameters.

Cell Lineage	Mean (μm)	Median (μm)	SD (μm)	Range (μm)
Neutrophils (n = 80)	12.15	12.23	1.04	11.00–13.06
Lymphocytes (n = 70)	9.44	9.53	1.36	8.38–10.17
Eosinophils (n = 28)	12.01	12.20	0.76	11.23–12.71
Monocytes (n = 30)	13.00	13.12	1.20	11.14–14.55
Erythrocytes (n = 134)	6.32	6.39	0.57	5.86–6.80

Mean values, median values, standard deviation, and overall ranges for the diameter of each observed cell type.

Fig 3

Individual cell diameters observed in neutrophils (n = 80), lymphocytes (n = 70), eosinophils (n = 28), monocytes (n = 30), and erythrocytes (n = 134).

Each color represents an individual bat with larger dots depicting the mean for each cell type per bat. The error bars display the mean and 95% confidence intervals for each cell type.

Lymphocytes

Lymphocytes were the dominant leukocyte type in 7.27% of individuals (n = 8/110) (Tables 1 and 2 and Fig 1). Lymphocytes displayed a high variability in size, shape, and color (Figs 2I–2P and 3 and Table 3). In general, lymphocytes were the smallest leukocyte type but varied in size, sometimes appearing similar in diameter to neutrophils (Table 3 and Fig 3). Most individuals had dimorphic or polymorphic lymphocyte populations. The cells were round to oval in shape, with a high nuclear to cytoplasmic ratio. Nucleus morphology was variable (round, oval, elliptical, or bean-shaped). A circular pale nucleolus was easily visible in a majority of larger lymphocytes. Binuclear lymphocytes were also observed in 2.72% of individuals (n = 3/110). These cells contained two clearly distinct nuclei of similar diameter (Fig 2N). A low number of reactive lymphocytes were regularly observed. Reactive lymphocytes were generally larger in size than non-reactive lymphocytes, showing distorted cytoplasmic and nuclear morphology with dark purple cytoplasmic edges (Fig 2M).

Unique large granular lymphocytes (LGLs) were observed and counted as part of the heterogenous lymphocyte population in 28.18% of individuals (n = 31/110). When observed, LGLs comprised 3.8 to 13% of the general lymphocyte population. These lymphocytes were generally larger and lighter in color than the other lymphocytes typically observed and often had an oval shape. The cytoplasm contained evenly distributed, reddish-pink, round-to-oval granules (Fig 2O and 2P). Granules were relatively large and visually striking but varied in size between individuals and were occasionally observed to have indistinct membranes.

Eosinophils

Eosinophils were present in abundances that ranged up to 25% of the leukocyte population present in the samples (Tables 1 and 2 and Fig 1). Eosinophils (Fig 2Q–2X) presented similar diameter to neutrophils (Figs 2I–2P and 3 and Table 3) but smaller than monocytes (Figs 2Y–2Ff and 3 and Table 3). Eosinophil nuclei generally had three to four lobes, although some were hypersegmented (>4 lobes). Low numbers of band eosinophils were observed in 15.45% of individuals (n = 17/110) (Fig 2Q and 2R). The granules were relatively large and usually stained orange to pink, although a low number of granules were grey in color. Granules were often seen throughout only half to three-quarters of the cytoplasm, rather than equally distributed throughout the entirety of the cytoplasm (Fig 2S). Additionally, granules often stained lightly, making the differentiation of neutrophils and eosinophils challenging.

Monocytes

Monocytes were observed to be highly variable in terms of overall cell size and color, both within and between individuals (Figs 2Y–2Ff and 3 and Table 3). These cells were generally observed to be larger than all other leukocytic cell types. Occasionally, monocytes were observed to be slightly smaller than neutrophils. Monocyte cytoplasm generally stained light to dark blue-grey with a classic “ground glass” textural appearance. Several monocytes were observed with round, dark purple cytoplasmic granules. The nuclei of mature monocytes were smooth and C-shaped, while immature monocytes (primarily promonocytes) showed an irregular and oval or round nucleus with a lumpy appearance. The chromatin patterns in most monocytes were often much smoother, finer, and lighter in color than that of neutrophils. However, chromatin patterns appeared to be affected by staining, as even mildly overstained smears showed monocytes with clumped chromatin patterns and slightly darkened, blue cytoplasmic borders. Low numbers of activated monocytes were observed throughout the sample set. Both vacuolated and non-vacuolated monocytes were observed, with vacuoles of highly variable sizes present in 58.18% of individuals (n = 64/110) (Fig 2Y–2Aa and 2Dd). 50.9% of individuals (n = 56/110) displayed both vacuolated and non-vacuolated monocytes.

Basophils

Consistent with previous reports of healthy P. alecto presenting low numbers of basophils, these cells were not observed in the smears examined [21, 22].

Erythrocytes

Erythrocytes stained with DipQuik generally displayed a normochromic coloration ranging from light to dark blue. Mild anisocytosis and polychromasia was commonly observed, and spherocytes, and cells with low central pallor were observed throughout the sample set (Fig 2Kk and 2Ll). Echinocytes (Fig 2Kk) were observed in 10% of individuals (n = 11/110) throughout our sample set. Dacrocytes (Fig 2Mm) were present in 48.1% individuals (n = 53/100). These cells were significantly smaller in diameter than all leukocytes observed (Table 3 and Fig 3).

Platelets

Individual platelets were observed to be small and purple with a blurred, spiky plasma membrane; or as small, light grey vesicles containing small, round, dark purple granules. Platelets were observed either spread across smears or occasionally clinging to the sides of leukocytes. Platelet clumps were considered to be aggregations of multiple platelets and were observed in 59.9% of individuals (n = 56/110) (Fig 2Hh and 2Ii).

Parasites

Potential hemoparasites were observed in the blood samples from one bat (Fig 4). Intracellular corpuscles consistent with the appearance of intraerythrocytic gametocytes (parasitic precursor cells) were observed in over 50 erythrocytes. These potential parasites varied in size, and a limited number of extracellular stages of the organism were also observed during the differential (Fig 4D). Generally, all stages of the organism showed distinguishable membranes containing mottled, granule-like components which often resembled beads on a string. Many of the potential intraerythrocytic parasites also showed one or two large inclusions (Fig 4C and 4D). An eosinophil count of 0 was observed on the differential cell count of this individual.

Fig 4

Erythrocytes with signs of infection by possible Hepatocystis hemoparasites observed in a single individual captured at Redcliffe in December 2018.

Images taken at 1000x magnification.

Discussion

Cell morphology

By conducting an extensive manual review of blood smears collected from 134 adult P. alecto, we were able to identify several trends within each observed leukocyte type. Although granular neutrophils were observed in our samples, the granulation was likely not pathologically significant. Of the ten individuals in which this morphology was observed, two bats had evidence of lesions consistent with frostbite, and another had evidence of aural dermatitis. The other seven individuals had no noted abnormalities or injuries. Healthy neutrophils contain low numbers of primary granules, and although these granules typically do not stain vividly enough to be seen with light microscopy, the low numbers and even distribution of the granules observed in our samples is not suggestive of toxic granulation [23]. The variation in neutrophil lobulation observed in our sample set is consistent with the appearance of polymorphonuclear neutrophils (PMNs) in other bat species [24].

A low number of reactive lymphocytes were regularly observed, but this is likely not pathologically significant, as low numbers of reactive lymphocytes are regularly observed in blood smears from clinically healthy small mammals [23] (Fig 2M). Similarly, the monocytes observed in our samples were consistent with typical findings in other mammals. Monocytes were generally observed to be larger than all other leukocytic cell types and many contained vacuoles of varying sizes, which is consistent with observations made in other mammalian species [25]. Low numbers of activated monocytes were observed throughout the sample set, which is also typical for mammals [23].

Echinocytes (erythrocytes with crenated edges, Fig 2Kk) and dacrocytes (tear drop-shaped erythrocytes, Fig 2Mm) are two erythrocyte anomalies consistently observed in our samples which are most likely artifacts of sample collection or slide creation. Echinocyte formation in particular is frequently associated with prolonged smear storage time [26]. Given the low numbers of anomalous cells observed in individual smears and the possibility that the process of smear creation disrupted the morphology of the cells, these observations are likely not significant. The platelet clumps observed on our slides were also unlikely to be significant, as platelet clumps in blood smears are generally artifacts of venipuncture sample collection [27].

Neutrophils were observed to be the dominant cell type in the majority of samples, with lymphocytes rarely appearing as the dominant cell type on a given slide. Previous research has demonstrated that neutrophil dominance is a typical finding in many bat species, including clinically normal P. alecto [21, 22, 28]. Our results are also consistent with findings of neutrophil dominance in other flying fox species, including the Rodriquez Island flying fox (Pteropus rodricensis), Indian flying fox (P. giganteus), Christmas Island flying fox (P. melanotus natalis), and the grey-headed flying fox (P. poliocephalus). However, the island flying fox (P. hypomelanus) and Malaysian flying fox (P. vampyrus) were observed to be lymphocyte-dominant [28-31].

No basophils were observed in our sample set, which is consistent with the low numbers of basophils typically reported in mammals, including other flying fox species [22, 24, 28, 30, 31]. The average numbers of eosinophils and monocytes observed on the smears in our study were also consistent with what has been reported previously [21, 22]. The low numbers of reactive neutrophils, monocytes, and lymphocytes observed across our sample set are consistent with findings in other mammalian species [23].

The large granular lymphocytes observed in 31 individuals (n = 31/110; none had observed injuries) were present across numerous samples from both sites. The distinctive morphological appearance we observed is consistent with descriptions of similar cells observed and described in multiple species of Neotropical bats [32]. We propose that these cells are natural killer (NK) cells or cytotoxic T cells, both of which have the appearance of normal lymphocytes but contain cytoplasmic granules [23]. However, additional cytochemical experiments are needed to confirm the identity of these cells. The apparent absence of reports of these cell types in other studies might result from the use of automated cell counts, which may not have differentiated these cells from other lymphocytes [21, 22]. By using a manual review of our smears, we were able to identify this consistently seen morphological anomaly as a noteworthy feature.

The intraerythrocytic hemoparasites observed in a single adult female from Redcliffe have a morphology that is consistent with Hepatocystis species. Although the appearance is generally consistent with reported Hepatocystis in other Pteropus species, molecular diagnostics would be needed to confirm taxonomic identity analysis that was not possible due to lack of appropriate samples [33]. This individual had an eosinophil count of zero; however, active endoparasitic infections are not always associated with eosinophilia at the time of infection [34]. Primary exposure to a parasitic organism generally results in delayed eosinophilia, which may manifest only after the parasitic organism dies, whereas subsequent exposures trigger intense, dramatic eosinophilia [23]. In our sample set, extremely elevated eosinophil counts (counts of 26% and 28% of observed cells, respectively), which may be evidence of recent parasitic pressure, were observed in one male and one female from Redcliffe captured in July and December 2019, respectively.

Although this study provides information about hematology in P. alecto, there are several limitations that should be considered. Due to the use of manual differential counts rather than flow cytometry or another automated counting technique, we are unable to report the total white blood cell concentration. While we were unable to state cell concentrations, we could report the proportions of leukocytic cells observed in our differential counts. The expense of automated methods and the need for fresh blood samples preclude the use of flow cytometry and other automated methods for population scale studies such as this. For population-level analyses, the relative proportions of leukocytic cells allows comparison of population hematological characteristics across large extents of space and time. In future studies, with larger sample sizes, such methods could be used to analyze temporal and more extensive spatial trends among reservoir host populations [35]. A previous study on this topic found seasonal differences in the neutrophil and lymphocyte counts which varied between males and females [21]. Finally, we focused on the hematology of adult members of this species, meaning that we were unable to report on the normal hematology of juvenile and sub-adult bats.

Conclusion

We report the first qualitative assessment of the leukocytes, erythrocytes, and platelets of clinically healthy adult P. alecto. Our study also provides a set of values comparing the relative abundance of different leukocytes in adult P. alecto. Considering the time span of our sampling (2 years), our results are likely to capture the natural variability of cell populations in healthy, wild individuals. By examining our hematological data both qualitatively and quantitatively, we provide both an in-depth characterization of the normal hematological parameters for P. alecto and examine the variations in leukocyte ranges between males and females.

Due to their important ecological role as pollinators and seed dispersers, Pteropodidae bats are of significant interest to conservation efforts. Gaining a better understanding of the normal hematological profile of P. alecto, will facilitate efforts to monitor population health, which could contribute to the identification of populations under stressful conditions. Research of this nature will not only advance studies of P. alecto hematology and population health, but also inform studies of related species. A number of other Pteropus species are of immediate interest to the study of disease spillover, including Pteropus medius, Pteropus lylei, and Pteropus vampyrus, which have all been identified as Nipah virus reservoirs [11]. Reservoir health is intricately intertwined with the spillover of zoonotic disease. By gaining a better understanding of normal hematology in Pteropus bats, we can better monitor wildlife populations with the potential to shed zoonotic viruses.

Sampling effort across sites, sexes, and years.

(DOCX)

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13 Dec 2021

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23 Feb 2022

Dear Dr. Becker,

RE: Revisions on manuscript PONE-D-21-35366 'Morphological and quantitative analysis of hemocytes in free-living Australian black flying foxes (Pteropus alecto)'

We appreciate the time and constructive critiques and comments from the reviewers and yourself. We have revised the manuscript accordingly.

Please, see below the specific responses to each of the comments. We present our responses, preceded by an “R”.

Editor comments:

L80: Is there a reference the authors can cite for the ordinal body condition score?

R: Per the recommendation of reviewer # 1, we have removed the body condition score statement from our methods.

L124: Can the authors remind the reader what fraction of individual blood smears were from adults?

R: All the individual blood smears used in this study came from adults. We provide this information in lines 132-134 as follows: “Considering the low number of individual bats from which good quality smears were obtained, we opted to summarize the data providing inter-quartile ranges per sex and using only adult individuals (n=134/134).”.

L128: Note that the log transformation is not appropriate for proportion data, as proportion data are bound between 0-1 (see Warton & Hui 2011 Ecology for a full discussion). I suggest the authors consider logit transforming proportions, which also facilitates back-transforming the proportions to the true 0-1 boundary of the data.

R: We agree that the logit transformation suggested would have been appropriate to handle the data in a formal statistical analysis. In the specific case of the data presented, logit transformation would require extra manipulation of the data to account for zeros in the dataset, which were common for cell populations present in low abundances. We transformed the data only for visualization purposes and allow readers to back-transform the data if needed in a more intuitive and easier process than if it were done from a logit transformation. To make sure that values of zero in the transformed data presented do not confuse the readers, we have added the following phrasing in the figure legend for clarification in back-transformation: “Values of cell populations present in low abundances were log-transformed as ln(x+1) for visualization purposes”.

L159: Perhaps clarify here that “individuals” = “adult bats”.

R: We have made this change. Please see lines 172-173 “Neutrophils were the dominant leukocyte type, comprising over 50% of leukocytes in 92.72% of adult bats (n=102/110).”

L160: Was a statistical test used (here and elsewhere) to compare leukocyte proportions between sexes and/or healthy/unhealthy bats? Or is this statement based only on data visualizations and raw summary statistics? I would suggest the authors consider the former as a more definitive way to make comparisons among groups.

R: Since the focus of our work was to present a reference of images and values for hematology of wild black flying foxes, we did not perform a formal statistical analysis, but rather focused on presenting the ranges of values. Though, we agree with the comment regarding formal comparisons. In this sense we consider that data on absolute numbers of cells which are not available for our dataset are needed to draw robust statistical comparisons. Acknowledging this caveat of our dataset, we adopted a conservative approach by presenting the dataset in a more descriptive way that could serve as baseline for future studies exploring the drivers of changes in leukocyte populations.

L275: Intraerythrocytic gametocytes of which parasites? It would help to identify the parasites to genus if possible, if not broader taxonomic level like order (e.g., haemosporidia).

R: Unfortunately, no sample is available on which to run a PCR or similar test to confirm taxonomic identification of the potential hemoparasites. However, we have included a phrase referring to previous studies in which hemoparasites have been described. Please see lines 369-372 “Although the appearance is generally consistent with reported Hepatocystis in other Pteropus species, molecular diagnostic would be needed to confirm taxonomic identity [33]), analysis that was not possible due to lack of appropriate samples.”

General editorial comments:

3. Thank you for stating the following in the Acknowledgments Section of your manuscript: We acknowledge the Kabi Kabi and Yuggera Ugarapul people, who are the Traditional Custodians of the land upon which this work was conducted. This study was funded by the DARPA PREEMPT program Cooperative Agreement # D18AC00031, and the U.S. National Science Foundation (DEB-1716698). DH and BH were funded by the Montana State University Undergraduate Scholars Program. BH was also funded by the Montana State University McNairs Scholar Program (Grant #P217A130148). RKP was also funded by the USDA National Institute of Food and Agriculture (Hatch project 1015891). AJP was supported by an ARC DECRA fellowship (DE190100710). KR acknowledges the support of The University of Sydney Institute for Infectious Diseases. Special thanks to Dr. Charlotte Hollinger, Dr. Dee McAloose, Dr. Siobhan Egan, Lauren Warner, Amelia Graves, and Lindsay Lee.

We note that you have provided funding information that is not currently declared in your Funding Statement. However, funding information should not appear in the Acknowledgments section or other areas of your manuscript. We will only publish funding information present in the Funding Statement section of the online submission form. Please remove any funding-related text from the manuscript and let us know how you would like to update your Funding Statement.

R: We have removed the funding-related text from the Acknowledgments.

Currently, your Funding Statement reads as follows: RKP was funded by the DARPA PREEMPT program (https://www.preemptproject.org) Cooperative Agreement # D18AC00031, and the U.S. National Science Foundation (https://www.nsf.gov) (DEB-1716698). DH and BEH were funded by the Montana State University Undergraduate Scholars Program (https://www.montana.edu/usp/). BEH was also funded by the Montana State University McNairs Scholar Program (https://www.montana.edu/mcnair/) (Grant #P217A130148). RKP was also funded by the USDA National Institute of Food and Agriculture (https://nifa.usda.gov) (Hatch project 1015891). AJP was supported by an ARC DECRA fellowship (https://www.arc.gov.au/grants/discovery-program/discovery-early-career-researcher-award-decra) (DE190100710).The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Please include your amended statements within your cover letter; we will change the online submission form on your behalf.

R: Our amended funding statement reads as follows: “RKP was funded by the DARPA PREEMPT program (https://www.preemptproject.org) Cooperative Agreement # D18AC00031, and the U.S. National Science Foundation (https://www.nsf.gov) (DEB-1716698). DH and BEH were funded by the Montana State University Undergraduate Scholars Program (https://www.montana.edu/usp/). BEH was also funded by the Montana State University McNairs Scholar Program (https://www.montana.edu/mcnair/) (Grant #P217A130148). RKP was also funded by the USDA National Institute of Food and Agriculture (https://nifa.usda.gov) (Hatch project 1015891). AJP was supported by an ARC DECRA fellowship (https://www.arc.gov.au/grants/discovery-program/discovery-early-career-researcher-award-decra) (DE190100710). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. The content of the information does not necessarily reflect the position or the policy of the U.S. government, and no official endorsement should be inferred.”

4. We note that you have stated that you will provide repository information for your data at acceptance. Should your manuscript be accepted for publication, we will hold it until you provide the relevant accession numbers or DOIs necessary to access your data. If you wish to make changes to your Data Availability statement, please describe these changes in your cover letter and we will update your Data Availability statement to reflect the information you provide.

R: There are no changes we wish to make to the Data Availability statement.

5. One of the noted authors is a group or consortium Bat One Health. In addition to naming the author group, please list the individual authors and affiliations within this group in the acknowledgments section of your manuscript. Please also indicate clearly a lead author for this group along with a contact email address.

R: The acknowledgments section of the manuscript has been updated to include the individual authors and affiliations within the BatOneHealth group. Please see lines 432-456 of the revised manuscript. The lead authors for the BatOneHealth group are Raina K. Plowright (raina.plowright@montana.edu) and Alison J. Peel (a.peel@griffith.edu.au).

6. We note that Figures 2 and 3 in your submission contain copyrighted images.

R: All images included in Figures 2 and 3 were taken by the authors for the purposes of this work.

Reviewer 1:

Broad edits/comments:

It seems like you took the natural log of eosinophils and monocytes. Perhaps, I missed it but could you provide a short statement on why you did this for these two cell types and not neutrophils and lymphocytes.

R: We have added this statement. Please see lines 137-139 “Histograms were used to assess the normality of the data, and natural log-transformations in the form ln(x+1) were applied to the monocyte and eosinophil counts to facilitate visualizations by adjusting for skewness.”

You mention that you calculate body condition scores in your methods section (line 80). I suggest either include (1) an analysis that explicitly looks at the relationship between body condition and cell counts, (2) create a table that summarizes counts from bats that are in poor (i.e. below a threshold value) or good (i.e. above a threshold value), or (3) that you remove this statement from your methods.

R: We removed this statement from our methods. The body condition scores are based on a commonly used field assessment. We used the score to roughly filter individuals that may present clinical signs of unhealthy condition. We consider the scoring too coarse/inaccurate to include in a formal analysis and it may mislead conclusions.

Minor edits:

Author list: “Bat One Health” is listed as an author with no affiliation details on the main manuscript document. Is this a mistake?

R: The full individual author and affiliation details can now be found in the acknowledgements section. Please see lines 432-456.

Line 95: As they’re anesthetized it likely doesn’t matter as much, but could you state when the blood sample was taken? For example, prior to or after the PIT tag insertion.

R: The blood samples were taken prior to PIT tag insertion. We have added this information to the methods section. Please see lines 96-97, “Prior to PIT tag insertion, a maximum of 2.5 mL of blood was drawn from the cephalic or uropatagial vein, with samples not exceeding 0.6% of body mass.”

Line 122: Remove the extra space

R: We have made this change. Please see line 132.

Line 275: Perhaps state what a gametocyte is? (e.g. precursor cells of a parasite)

R: We have defined gametocytes as “parasitic precursor cells” in lines 297-300. “Potential hemoparasites were observed in the blood samples from one bat (Fig. 3). Intracellular corpuscles consistent with the appearance of intraerythrocytic gametocytes (parasitic precursor cells) were observed in over 50 erythrocytes”

Figure 3 caption: Could you put in the caption which hemoparasite(s) you believe these to be?

R: We have updated the caption to include this information. Please see lines 309-310 “Fig 3: Erythrocytes with signs of infection by possible Hepatocystis hemoparasites observed in a single individual captured at Redcliffe in December 2018. Images taken at 1000x magnification.”

Reviewer 2:

1. through the entire manuscript please replace hemocytes with white blood cells or leukocytes, as hemocytes refers to the cells of invertebrates

R: We have made this change throughout the manuscript.

2. in the abstract it is written “…leukocyte morphological traits that are relevant for future research on bat health, especially in context of viral spillover and emergence”. I could not find these in the discussion. Would it be possible to rephrase this sentence, present and focus more the specific results?

R: Considering this comment, we have rephrased the sentence. See lines 15-16. “...leukocyte morphological traits that are relevant for future research on bat health, including unique large granular lymphocytes.”

3. introduction – “bat-borne pathogens”: are all these serious threat to human health, or some of them only? Again I think being more specific on viruses and/or intracellular pathogens, would help also to decrease the negative distinction of bats received lately due to their reservoir competence.

R: We appreciate this comment and agree with the reviewer’s point. Accordingly, we have changed the language, see lines 22-23, “Several bat-borne viruses are of human health concern when cross-species transmission events, or spillovers, occur." In addition, we added information to the introduction regarding the ecological importance of the species in lines 48-53: “One species of interest is Pteropus Alecto (black flying fox) which plays important ecological roles as pollinators and seed dispersers, and are endemic to Australia, Indonesia, and Papua New Guinea [15]. Despite the role of P. alecto as reservoirs of Hendra virus, relatively few studies have tried to characterize the physiological condition of clinically “healthy” individuals to serve as baseline for health assessments.”

4. line 34 – “medical” history

R: Comment included in line 33: “…largely due to a lack of suitable metrics in wild animals whose medical history is unknown”.

5. lines 50 – 60: other drawbacks of automated methods are logistics (e.g. remote field conditions) and volume requirements (e.g. smaller sized species). These could be included in this section too.

R: We agree with this comment and have included it in the text. Please see lines 54-58. “This approach, although highly efficient in terms of human hours and samples processed, can be impractical due to the logistics of using cytometers in remote field conditions, and the higher sample volume requirements for automated counting relative to slide preparation. Importantly, automated cell counters can sometimes present errors associated with variability in cell types among species.”

6. The authors report and discuss the size of each cell type relative to others, which is indeed helpful when analyzing blood smears. However, I was wondering whether it would be possible to measure the size and thickness of each cell type and report these too?

R: We have analyzed a subset of our samples to report the diameter of each cell type (see lines 117-123). We have added an additional table and figure to present intra and inter-individual variation of cell sizes. Please see Table 3: Cell Diameters, and figure 3.

7. figure 1 – please rearrange the order as differs between the figure and the order described in the legend

R: Figure 1 has been rearranged in the following order: neutrophils, lymphocytes, eosinophils, and monocytes. The legend has been updated accordingly.

8. figure 1 and figure 2 – I would suggest to keep the logical order from the manuscript: neutrophils, lymphocytes, eosinophils and monocytes.

R: These changes have been made. Please see Figures 1 and 2.

9. references - through the manuscript there were few sentences where I missed references supporting the statements. Could you please include these, as this would help the reader? See lines 320 (on neutrophil dominance in mammals); lines 356-364 (reference 19 is not the right one, please change it; LPS induced systemic inflammation causes neutrophilia in various bat species, including in Rousettus aegyptiacus; maybe some of these papers can be included here); lines 377-379 (indeed differential counts can allow comparison across larger scales. Reference missing – see Becker et al. 2019 Integr Comp Biol on common vampires).

R: The references in line 320 of the original manuscript were specific to bat species rather than mammals generally, so we have changed the language to reflect this. Please see lines 342-345: “Previous research has demonstrated that neutrophil dominance is a typical finding in many bat species, including clinically normal P. alecto (22,23,28).”. The change in references in lines 356-364 has been made, please see reference 21 in line 382-383 of the revised version. We have also included the recommended reference for lines401-405as reference 38.

10. I was surprised that the authors did not perform total white blood cell counts using standardized method with hemocytometer…It requires only small amount of blood and a field microscope. Why was this omitted? Although the differential counts alone have their values as discussed in this paper, still would be great if more studies would aim for total leukocyte counts too and not only via automated methods.

R: We agree that this would have been a valuable analysis that would have contributed to a more quantitative approach (see our reply about statistical approach recommended by the editor), unfortunately this information was not collected due to time constraints for the field team to perform the analysis. To keep the fieldwork safe for the team, we tried to keep field days under 12 hours. We will consider field cell counts during future smaller studies.

11. Besides the value of such study for disease research, I think also has importance for conservation (e.g. identifying anthropogenic stressors, species of conservation interest) – this aspect should be mentioned in the conclusion section.

R: We appreciate the comment. We have updated both the introduction and the conclusion to include this information. Please see lines 46-53 “As contact between flying fox populations, domestic animals, and humans increases, so does the risk of transmission of zoonotic infectious pathogens (3,4,6,7). One species of interest is the black flying fox (Pteropus alecto). Bats of this species play an important ecological role as pollinators and seed dispersers, and are endemic to Australia, Indonesia, and Papua New Guinea (15). Despite the role of black flying foxes as reservoirs of Hendra virus, relatively few studies have tried to characterize the physiological condition of clinically “healthy” individuals to serve as baseline for health assessments.”

And lines 417-420:

“Due to their important ecological role as pollinators and seed dispersers, Pteropus bats are of significant interest to conservation efforts. Gaining a better understanding of the normal hematological profile of P. alecto, will facilitate efforts to monitor population health, which could contribute to the identification of populations under stress.”

Thank you for your time and valuable comments.

Sincerely,

Dale Hansen, Brooklin E. Hunt, Caylee A. Falvo, Manuel Ruiz-Aravena, Maureen K. Kessler, Jane Hall, Paul Thompson, Karrie Rose, Devin N. Jones, Tamika J. Lunn, Adrienne S. Dale, Alison J. Peel, Raina K. Plowright

Submitted filename: Response to Reviewers.docx

Click here for additional data file.

14 Mar 2022

28 Apr 2022

Dear Dr. Becker,

RE: Revisions on manuscript PONE-D-21-35366 'Morphological and quantitative analysis of lymphocytes in free-living Australian black flying foxes (Pteropus alecto)'

We greatly appreciate the time you have taken to review this manuscript. We agree with the edits you have suggested, and after considering the two courses of revision you laid out in your review we have opted to keep the descriptive focus of this paper and remove any inferential language. This had led to changes in the results and discussion sections. Please see lines 176, 242, 272, 282, and 412 for the revisions made in these sections. We have additionally revised our methods section to clarify that we have not performed any formal statistical analyses. Please see lines 141-143.

As a result of the changes made to the discussion section, we are no longer including the following three references:

35. Pacioni C, Robertson ID, Maxwell M, van Weenen J, Wayne AF. Hematologic Characteristics of the woylie (Bettongia penicillata ogilbyi). J Wildl Dis. 2013;49: 816–30.

36. Zuk M, McKean KA. Sex differences in parasite infections: Patterns and processes. Int J Parasitol. 1996;26: 1009–24.

37. Christe P, Glaizot O, Evanno G, Bruyndonckx N, Devevey G, Yannic G, et al. Host sex and ectoparasites choice: preference for, and higher survival on female hosts. J Anim Ecol. 2007;76: 703–10.

To the best of our knowledge, the rest of the reference list is complete and correct, and we cite no retracted articles.

Thank you for your time and valuable comments.

Sincerely,

Dale Hansen, Brooklin E. Hunt, Caylee A. Falvo, Manuel Ruiz-Aravena, Maureen K. Kessler, Jane Hall, Paul Thompson, Karrie Rose, Devin N. Jones, Tamika J. Lunn, Adrienne S. Dale, Alison J. Peel, Raina K. Plowright

Submitted filename: Response to Reviewers.docx

Click here for additional data file.

3 May 2022

Morphological and quantitative analysis of leukocytes in free-living Australian black flying foxes (Pteropus alecto)

PONE-D-21-35366R2

Dear Dr. Hansen,

We’re pleased to inform you that your manuscript has been judged scientifically suitable for publication and will be formally accepted for publication once it meets all outstanding technical requirements.

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PLOS ONE

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Reviewers' comments:

10 May 2022

PONE-D-21-35366R2

Morphological and quantitative analysis of leukocytes in free-living Australian black flying foxes (Pteropus alecto)

Dear Dr. Hansen:

I'm pleased to inform you that your manuscript has been deemed suitable for publication in PLOS ONE. Congratulations! Your manuscript is now with our production department.

If your institution or institutions have a press office, please let them know about your upcoming paper now to help maximize its impact. If they'll be preparing press materials, please inform our press team within the next 48 hours. Your manuscript will remain under strict press embargo until 2 pm Eastern Time on the date of publication. For more information please contact onepress@plos.org.

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