PROOF ONLYREPRODUCTION

© 2017 Society for Reproduction and Fertility DOI: 10.1530/REP-16-0578ISSN 1470–1626 (paper) 1741–7899 (online) Online version via www.reproduction-online.org

ReseaRch

Isolation of luteal endothelial cells and functional interactions with T lymphocytes

S S Walusimbi1, L M Wetzel1, D H Townson2 and J L Pate1

1Department of Animal Science, Center for Reproductive Biology and Health, Pennsylvania State University, University Park, Pennsylvania, USA and 2Department of Animal and Veterinary Sciences, University of Vermont, Burlington, Vermont, USA

Correspondence should be addressed to J L Pate; Email: jlp36@psu.edu

abstract

The objectives of this study were to optimize the isolation of luteal endothelial cells (Lec) and examine their functional interactions with autologous T lymphocytes. analysis by flow cytometry showed that the purity of Lec isolated by filtration was nearly 90% as indicated by Bandeiraea simplicifolia (Bs)-1 lectin binding. Lec expressed mRNa for progesterone receptor (PGR), prostaglandin receptors (PTGFR, PTGeR2 and 4, and PTGIR), tumor necrosis factor receptors (TNFRsF1a&B) and interleukin (IL) 1B receptors (IL1R1&2). Lec were pretreated with either vehicle, progesterone (P4; 0–20 µM), prostaglandin (PG) e2 or PGF2α (0–0.2 µM), and further treated with or without TNF and IL1B (50 ng/mL each). Lec were then incubated with autologous T lymphocytes in an adhesion assay. Fewer lymphocytes adhered to Lec after exposure to high compared to low P4 concentrations (cubic response; P < 0.05). In contrast, 0.2 µM PGe2 and PGF2α each increased T lymphocyte adhesion in the absence of cytokines (P < 0.05). Lec induced IL2 receptor alpha (cD25) expression and proliferation of T lymphocytes. In conclusion, filtration is an effective way of isolating large numbers of viable Lec. It is proposed that PGs and P4 modulate the ability of endothelial cells to bind T lymphocytes, potentially regulating extravasation, and that Lec activate T lymphocytes migrating into or resident in the cL.Reproduction (2017) 153 519–533

Introduction

The corpus luteum (CL) is a transient endocrine gland with a high vascular density similar to that observed in solid tumors (Bruce & Moor 1976, Zheng et al. 1993). The vasculature constitutes about 30% of the total volume of the bovine CL (Singh et al. 1997). Although estimates of cell numbers in tissue sections indicate that endothelial cells are about 50% of all cells in ovine (Rodgers et al. 1984, Azmi & Bongso 1985) and bovine CL (O’Shea  et  al. 1989), endothelial cells comprised 88.4% of total dispersed ovine luteal cells (Rodgers & O’Shea 1982). Additionally, based upon in vitro studies, luteal endothelial cells (LEC) are described as exhibiting five morphological phenotypes (Spanel-Borowski 2011), with each contributing to the development and function of the CL (Shirasuna et al. 2012a).

Although LEC are abundant in the CL, current isolation methods recover relatively few of these cells to conduct extensive experiments in vitro. Density gradient centrifugation (Rodgers & O’Shea 1982, Spanel-Borowski & van der Bosch 1990, 1993, Plendl  et  al. 1996a,b), fluorescence activated cell sorting (FACS; Voyta  et  al. 1984, van Beijnum et al. 2008), and lectins or antibodies ligated to magnetic beads (MACS; Drake & Loke 1991, Christenson & Stouffer 1996, Klipper  et  al. 2004,

Zannoni et al. 2007) are among the methods used, but the proportion of LEC recovered is far less than the estimates of 50% or 88% LEC reported in the above studies. Poor recovery of LEC for primary culture has led to alternative approaches using immortalized cells (Korzekwa  et  al. 2011) and the use of passaged LEC (Cavicchio et al. 2002, Cherry et al. 2008), which may not entirely preserve the characteristics of LEC. Therefore, new methods designed to maximize the recovery of primary endothelial cells from dispersed luteal cells would be insightful.

In addition to delivering nutrients to support steroid synthesis of the steroidogenic cells (Wiltbank  et  al. 1988), the endothelium of the CL regulates the trafficking of immune cells into the parenchyma. Similar to endothelial cells found in other tissues, LEC express adhesion molecules such as selectin P (Rohm et al. 1990), chemokines such as chemokine (C–C motif) ligand 2 (CCL2) and IL8 (Townson et al. 2002, Shirasuna et al. 2012b), which bind, arrest and guide the extravasation process of leukocytes into tissues (Newman  et  al. 1990, Bochner  et  al. 1991, Walter & Issekutz 1997, Woodfin  et  al. 2007). Endothelial cells also express class II major histocompatibility molecules (MHC II) and costimulatory molecules, such as lymphocyte

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function-associated antigen 3 (LFA3/CD58), cluster of differentiation (CD)40 and CD80 (Ashida  et  al. 1981, Yellin et al. 1995, Karmann et al. 1996, Cannon et al. 2007a,b), which are involved in T lymphocyte activation (Hughes et al. 1990). Activated T lymphocytes present in the bovine CL (Penny et al. 1999, Bauer et al. 2001, Townson et al. 2002, Poole & Pate 2012) can modulate steroidogenesis (Nothnick & Pate 1990, Fairchild & Pate 1991, Benyo & Pate 1992, Townson & Pate 1996, Castro et al. 1998) and are proposed to augment luteal regression (Pate 1995 rev.; Bukovský  et  al. 1995). Immune cells that migrate into the CL play roles in luteal development, homeostasis and luteolysis (Pate 2012, Walusimbi & Pate 2013).

The number and type of immune cells in the CL is dynamic and changes throughout the estrous cycle (Walusimbi & Pate 2013). For instance, in the bovine CL there are few T lymphocytes present during luteal development, but their numbers increase in a fully functional CL (days 10–14), and remain elevated throughout the remainder of the luteal phase (Lobel & Levy 1968, Penny  et  al. 1999, Bauer  et  al. 2001, Townson  et  al. 2002). However, the mechanisms orchestrating these temporal fluctuations of immune cells are not fully understood. Additionally, there is little known about the cellular mechanisms that augment T lymphocyte adherence, extravasation, and activation within the CL, activities known to depend upon the relative state of endothelial cell and T lymphocyte activation (Spanel-Borowski 2011, Walusimbi & Pate 2013). The CL synthesizes vast amounts of steroids, prostaglandins (PG; Milvae & Hansel 1983, Milvae 1986, Hayashi  et  al. 2003, Waclawik  et  al. 2008, Weems  et  al. 2012), cytokines (Shaw & Britt 1995, Petroff et al. 1999, Penny et al. 1999, Cavicchio et al. 2002, Neuvians et al. 2004) and chemokines (Townson & Liptak 2003). All of these molecules regulate the expression of adhesion molecules on endothelial cells and the adhesion of immune cells to endothelial cells in vitro (Winkler et al. 1997, Simoncini et al. 2004). In particular, the direct effect of cytokines and chemokines on the process of immune cell recruitment into lymphoid and nonlymphoid tissues is well established, but that of P4 and PG is still unclear. In cattle, the number of immune cells increases during luteal regression as P4 synthesis declines (Penny  et  al. 1999, Shirasuna  et  al. 2012a). With regard to PG, the synthesis, transport and signaling pathways of intraluteal PGE2 are more prominent during luteal development and maintenance, whereas the PGF2α system becomes activated during luteolysis (Arosh et al. 2004). Based upon the patterns of P4 and PG synthesis, secretion and action, it is conceivable that luteal P4 and PG modulate endothelial cell and T lymphocyte activation and interaction within the CL. In the current study, P4, PG and cytokines were hypothesized to act directly on luteal endothelial cells to modulate adhesion of T lymphocytes. Additionally, a new

and innovative filtration method to isolate LEC was tested and compared to magnetic sorting to investigate the above functional interactions between LEC and T lymphocytes in the presence and absence of progesterone, prostaglandins and cytokines.

Materials and methods

Reagents and antibodies

Reagents used for polymerase chain reaction (PCR) were: TRIzol reagent (Invitrogen), iScript cDNA synthesis kit (Bio-Rad), DyNAmo cDNA synthesis kit (Thermo Fisher), Power SYBR Green PCR master mix (Applied Biosystems), SensiMix SYBR Green No-ROX kit (Bioline), RNase-free DNase I (Ambion) and QiAquick Gel Extraction Kit (QIAgen). Reagents used for cell culture were: Ham’s F-12 nutrient mixture; AIMV cell culture medium and RPMI 1640 medium (1×; without phenol red; Gibco, Invitrogen); carboxyfluorescein succinimidyl ester (CFSE; Sigma); insulin, transferrin and selenium (ITS; VWR); bovine serum albumin (BSA; Sigma); phorbol 12-myristate 13-acetate (PMA) and ionomycin calcium salt (Sigma); EGM-2 (EBM-2 plus SingleQuot kit of growth factors; Lonza); rbTNF (2279-BT-025; R&D Systems) and rbIL1B (RBOIL1BI; Pierce). Antibodies used for the isolation of T lymphocytes and endothelial cells were: CD2 (MUC2A; VMRD), anti sheep CD31 (AbD Serotec), anti mouse IgG2a, anti mouse IgG1 and IgG2a + b magnetic beads (Miltenyi). Antibodies used for T lymphocyte phenotyping were: δTCR-N12 (CACT61A; VMRD), CD8α (BAQ111A; VMRD), CD8β (BAT82A; VMRD), WC1 (IL-A29; VMRD), anti mouse IgM:PE (STAR81F; AbD Serotec) and anti mouse IgG1:PE (STAR81PE; AbD Serotec). Antibodies used for immunostaining were: anti human von Willebrand Factor (Dako) and anti-human VE-cadherin (AHP628Z; AbD Serotec). Calcein AM (Vybrant cell adhesion assay kit) was purchased from Invitrogen. Reagents used for western blot were: CelLytic MT Cell Lysis Reagent (Sigma-Aldrich), SuperSignal West Femto Maximum Sensitivity Substrate (Thermo Scientific), Pierce BCA Protein Assay Kit (Thermo Scientific), Restore PLUS Western Blot Stripping Buffer (Thermo Scientific), horseradish peroxidase-labeled anti rabbit secondary antibody (GE Healthcare), polyvinylidene difluoride membranes (Hybond-P; Amersham Pharmacia Biotech) and polyclonal rabbit anti mouse VCAM1 antibody (bs-0396R; Bioss).

Isolation of cells

Corpora lutea were collected transvaginally from cyclic Holstein cows following the procedure approved by the Institutional Animal Care and Use Committee of Pennsylvania State University. The CL were processed following procedures detailed by Pate (1993) with the following modifications: Collagenase-dispersed cells were filtered through a 50-μm pore size nylon microsieve (BioDesign Inc.) and the filtrate was serially centrifuged at 200, 128, 72 and 32 g, 10 min each time at 4°C, to pellet luteal steroidogenic cells (LSC). The supernatant after each centrifugation was collected in 50-mL polystyrene tubes (always kept on ice) and used for luteal

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endothelial cell (LEC) isolation by filtration and magnetic separation.

Isolation of LEC by filtration

The supernatant obtained by serial centrifugation of mixed luteal cells was further centrifuged at 300 g to obtain endothelial cells. The pellet was resuspended in 10 mL of phosphate buffered saline–2 mM EDTA (PBS–EDTA) and gently agitated for 5–10 min to allow separation of cell aggregates, then filtered sequentially through 15- and 10-μm pore size nylon microsieves into a glass beaker placed on ice. The filtrate was centrifuged at 300 g and the supernatant was discarded. Five to ten mL of cold, freshly prepared red blood cell lysis buffer (0.14 M NH4Cl, 10 mM KHCO3, 0.1 mM EDTA, pH 7.4) was added to the pellet for 5 min. The cells were subsequently diluted with excess EGM-2 cell culture medium and centrifuged at 300 g. The pellet was resuspended in EGM-2 and the number and viability of cells recovered was determined by flow cytometry (EasyCyte Plus, Millipore) after labeling with ViaCount dye (Guava ViaCount, Millipore). Forward and side scatter characteristics of cells obtained from the enriched LEC and mixed luteal cells (LC) were compared.

Isolation of LEC by magnetic separation

Endothelial cells were also isolated by magnetic cell separation on an autoMACS Pro Separator (Miltenyi, Germany) using anti-CD31 primary antibody (10 μg/mL) and general procedures described for isolation of immune cells (Ndiaye et al. 2008). The number and viability of cells were determined by flow cytometry after labeling with ViaCount dye.

Characterization of enriched endothelial cells

The characteristics of enriched LEC relative to LC isolated from midcycle (days 10–12) CL were evaluated by flow cytometric analysis. The relative purity of enriched LEC and efficiency of isolation were determined by binding of the endothelial cell-specific BS-1 lectin and quantification of StAR mRNA by

qPCR. Fresh cells (1 × 106) were incubated with 2 μg/mL of BS-1 lectin-FITC for 30 min. The cells were centrifuged three times at 300 g and resuspended in PBS-EDTA before flow cytometric analysis. Cells not labeled with BS-1 lectin-FITC were used as a control.

Endothelial cells were cultured for 6  days on coverslips in EGM-2 medium and examined for binding of BS-1 lectin, and expression of von Willebrand factor (vWF) and vascular endothelial (VE) cadherin. The cells were fixed with 4% paraformaldehyde for 15 min at room temperature, washed 2× with PBS and stained with either fluorescein (FITC)-conjugated BS-1 lectin (2 μg; Sigma) or with primary antibodies against vWF and VE-cadherin overnight at 4°C. Unstained cells were used as a control for BS-1 while cells stained with rabbit IgG were used as controls for vWF and VE-cadherin. After incubation with primary antibodies, the cells were washed 3× with PBS and further incubated with PBS + 5% goat serum to block nonspecific binding. Antirabbit secondary antibodies conjugated with FITC were added and the cells were incubated for 1 h at 4°C and subsequently washed 3× with PBS. The coverslips were mounted on glass slides with ProLong Gold Antifade Reagent (Invitrogen). Images of stained cells were acquired using a DP71 color camera mounted on a BX51 Olympus fluorescence microscope.

Messenger RNA (mRNA) was extracted with TRIzol reagent from LEC and mixed luteal cells that were stored at −80°C to determine the expression of steroidogenic acute regulatory protein (STAR). Ribosomal protein 19 (RPL19) was used as a reference gene. The concentration of RNA and the presence of contaminants (260/280 and 260/230 ratios) were determined by spectrophotometry (NanoDrop ND-1000; NanoDrop Technologies, Inc.). Total RNA was treated with RNase-free DNase1 to eliminate genomic DNA contamination and a total of 2 μg was reverse transcribed using random oligos and the iScript cDNA Synthesis Kit following the manufacturer’s protocol. Primers (Table 1) were validated by sequencing of products that were generated in a thermocycler (35 cycles), separated on a 1.5% agarose gel and extracted using QiAquick Gel Extraction Kit. The products were precipitated and purified using ethanol and used for generating standard curves for

Table 1 Primer sequences for qPCR of Bos taurus transcripts.

Gene

GenBank accession no.

Forward

Reverse

amplicon size (bp)

RPL19 NM_001040516.1 5′-ATCGATCGCCACATGTATCA-3′ 5′-GCGTGCTTCCTTGGTCTTAG-3′ 168TNFRSF1A NM_174674.2 5′-CAGTCTCAACCCTCCACTCCC-3′ 5′-CATCTGCCAACAGAGCTGGAA-3′ 199TNFRSF1B NM_001040490.2 5′-CAGGCTGTGTTTACCCCCTA-3′ 5′-TGTCCAAGGTCATGTTGCAT-3′ 142IL1R1 NM_001206735.1 5′-ATAAACCAGACGGCAGGACG-3′ 5′-GATGAGAGGACACGAGCGAG-3′ 211IL1R2 NM_001046210.2 5′-TTGTCAGCCTCTGGGCTACT-3′ 5′-CGAGTAAGCGAGTTCCCCTC-3′ 148PTGFR NM_181025.3 5′-CACAGACAAGGCAGGTCTCA-3′ 5′-CTCCACAACAGCGTCTGGTA-3′ 265PTGIR NM_001015622.2 5′-GGCAGGCGAGGACGAGGTTG-3′ 5′-TCCGAGAGTCGCCTTGGGCA-3′ 278PTGER2 NM_174588.2 5′-AATGCGTTCAGTCCTCTGCT-3′ 5′-TGCAAGAAGTGCCTTGTCAG-3′ 278PTGER4 NM_174589.2 5′-TCTCTGGTGGTGCTCATCTG-3′ 5′-GTCTTCCGCAGGAGGATGTA-3′ 173PGR AY656812 5′-GAGACCTCATCAAGGCAATTGG-3′ 5′-CACCATCCCTGCCAATATCTTG-3′ 227SELPa NM_174183.2 5′-AGTGTGTAGCTGTCCAGTGC-3′ 5′-ATCCCAGTTCCCCGTATCCA-3′ 359SELPb NM_174183.2 5′-TCGCATTAGTTGGGCCAGAG-3′ 5′-CACTTGCTCCCACCAGGAAT-3′ 207CCL2 NM_174006.2 5′-CTGCAACATGAAGGTCTCCG-3′ 5′-TTGGGGTCTGCACATAACTCC-3′ 243VCAM1 NM_174484.1 5′-TTGGATGGTGTTTGCAGTTTCT-3′ 5′-AGTCAGTGAAACAGAGTCACCAATCT-3′ 97STAR NM_174189.2 5′-GCATCCTCAAAGACCAGGAG-3′ 5′-CTTGACGTTGGGATTCCACT-3′ 194aPrimer pair used for PCR, first step of nested qPCR. bPrimer pair used with SELP PCR product for qPCR, second step of nested qPCR.

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quantification. Hot start real-time qPCR (7500 Fast Real-Time PCR System, Applied Biosystems) was used to quantify the steady-state amount of mRNA using Power SYBR Green PCR master mix and the following conditions: denaturing, 95°C for 30 s; annealing, 60°C for 45 s; extension, 72°C for 60 s; and extra elongation, 72°C for 5 min.

Assessment of LEC and T lymphocyte adhesion/interaction

Acknowledging several molecular pathways exist to mediate endothelial cell–T lymphocyte adhesion (Shimizu  et  al. 1991), the current study examined the vascular cell adhesion molecule 1 (VCAM1) pathway, principally based upon evidence that VCAM1 is expressed in bovine CL (Lehmann et al. 2000, Shirasuna et al. 2012b), and because cytokine-activated endothelial cells express VCAM1 in other tissues (Shimizu  et  al. 1991, Aziz & Wakefield 1996, Wolf  et  al. 2001, Ono  et  al. 2006). To evaluate LEC–T lymphocyte activation, interaction and then adhesion in vitro, (1) LEC expression of VCAM1 was assessed by immunoblotting; (2) LEC responsiveness to cytokine stimulation was tested by measuring genetic expression of cytokine/hormone receptors and adhesion molecules; (3) LEC-induced activation of T lymphocytes was studied using coculture, and measuring LEC-induced proliferation and phenotypic changes of T lymphocytes; and (4) LEC-induced T lymphocyte adhesion was assessed by fluorescence assay, as detailed below.

LEC expression of VCAM1

Luteal tissues collected on day 6 and days 10–12 of the estrous cycle, as well as 0, 0.5, 1, 2, 4, 8 and 12 h after an injection of PGF2α to induce luteal regression, were snap frozen in liquid nitrogen and stored at −80°C. Frozen tissues were homogenized and proteins extracted from tissue using CelLytic MT Cell Lysis Reagent following the manufacturer’s protocol.

Proteins were quantified using the Pierce BCA Protein Assay Kit. Protein samples (30 μg) were subjected to electrophoresis on a 12% SDS-polyacrylamide gel and the separated proteins were blotted onto polyvinylidene difluoride membranes. Specific proteins were identified using a polyclonal rabbit anti-mouse VCAM1 antibody. First, membranes were incubated with blocking buffer (20 mM Tris (pH 7.4), 150 mM NaCl, 5% nonfat dry milk, and 0.05% Tween 20; TBS-Tween) for 1 h at room temperature followed by an overnight (12–16 h) incubation at 4°C with anti-VCAM1 antibody at a final concentration of 10 μg/mL. Membranes were washed twice with TBS-Tween and incubated with horseradish peroxidase-labeled anti-rabbit secondary antibody at a dilution of 1:10,000 for 2 h at room temperature. The antigen–antibody complex was visualized using the enhanced chemiluminescence system following the manufacturer’s protocol. Membranes were exposed to SuperSignal West Femto Maximum Sensitivity Substrate and visualized using the Quantity One program and ChemiDoc (Bio-Rad). Membranes were stripped using Restore PLUS Western Blot Stripping Buffer, and beta actin (ACTB) was used as an internal control to verify the integrity of proteins in the samples.

LEC responsiveness to cytokine stimulation

Luteal endothelial cells isolated from midcycle CL via the filtration method described above were cultured in RPMI 1640 medium supplemented with 20% fetal bovine serum (FBS) and gentamicin in 6-well plates for 72 h to allow cells to adhere. Medium was changed to RPMI 1640 medium supplemented with 10% FBS for proliferation of cells for 24 h. Cells were cultured overnight in RPMI 1640 medium supplemented with gentamicin, insulin (5 μg/mL), transferrin (5 μg/mL) and selenium (5 ng/mL). On the following day, cells were treated with TNF (50 ng/mL) and IL1B (50 ng/mL) for 6 h, to determine the effects of cytokines on receptor mRNA concentrations for TNF (TNFRSF1A&B), IL1B

Figure 1 Characteristics of freshly enriched endothelial cells. The number (A; n = 3) of cells recovered after magnetic cell sorting (CD31+) and filtration. (B) Representative graphs of forward (cell size) and side (internal complexity) scatter characteristics of mixed luteal cells (before filtration) and enriched endothelial cells (after filtration). Regions (R1, R2 and R3) indicate distinct populations of cells based on size (forward scatter) and intracellular complexity (side scatter). (C) Proportion of viable cells recovered from CD31-mediated magnetic cell sorting and filtration (B; n = 3). (D) The proportion of BS-1+ cells (n = 6) and (E) steady-state abundance of STAR transcripts in cells retained by the 10-µm filter (retentate; n = 4) and enriched endothelial cells (filtrate). Means with different letters are significantly different (P < 0.05).

AB

C D EP

×

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(ILR1&2), PGF2α (PTGFR), PGE2 (PTGER2&4), PGI2 (PTGIR) and progesterone (PGR). In addition, cultured LEC treated with TNF (10 ng/mL for 24 h) and luteal tissue collected on day 6, days 10–12 of the estrous cycle, and during PGF2α-induced luteal regression, were used to determine concentrations of mRNA for selectin P (SELP), vascular cell adhesion molecule 1 (VCAM1) and chemokine (C–C motif) ligand 2 (CCL2).

Steady-state concentrations of mRNA were quantified using qPCR as described above, except using DyNAmo cDNA synthesis kit and SensiMix SYBR Green No-ROX. The annealing temperature of ILR2 was 58°C. Amplification of SELP mRNA was achieved using nested PCR. Endothelial cell cDNA was amplified by PCR using the first set of primers for 25 cycles. Ten (10) percent of the resulting PCR product was then used as the starting material for qPCR using the second set of primers under the conditions described above. The quantity of mRNA was evaluated in duplicate wells for each sample from at least three biological replicates. A relative standard curve method (1:10 serial dilutions) generated from the purified product was used to determine the relative amount of mRNA present in the samples. Samples without template (No RT) were used as negative controls for amplification.

LEC-induced proliferation and phenotypic changes of T lymphocytes (LEC–T lymphocyte interactions)

Jugular venous blood was collected from the same animals as the CL and peripheral blood mononuclear cells (PBMC) were obtained by centrifugation over Ficoll-Paque (GE Healthcare). T lymphocytes were isolated from PBMC by positive selection using an autoMACS Pro Separator (Miltenyi Biotech, Germany) and anti-CD2 and anti-γδ antibodies as described by Ndiaye et  al. (2008). The purity, evaluated as percentage of cells positive for CD3, was >95%. Cell numbers were determined as described above.

To determine if LEC induced T lymphocyte proliferation, T lymphocytes (12.5 × 106/mL) were labeled with carboxyfluorescein succinimidyl ester (CFSE; 1.75 μM). Briefly, CFSE-labeled T lymphocytes in AIMV medium were incubated at 37°C for 15 min and further incubated at room temperature for another 10 min. The cells were diluted 5-fold with medium containing 10% fetal calf serum and washed three times at 300 g for 10 min. The CFSE-labeled T lymphocytes (5 × 105) were cocultured with either LEC isolated by the filtration method seeded at 1 × 106 cells/well in 24-well plates or with LC (5 × 105/well). Proliferation was determined by CFSE dilution after 72 h.

To determine if LEC alter the activation status or phenotype of T lymphocytes, LEC or LC were cocultured with T lymphocytes. After 72 h, the T lymphocytes were harvested and expression of CD25, CD8α, CD8β, δTCR and WC1 were

A B

C D

E F

Figure 2 Representative images of enriched and cultured endothelial cells. (A) Freshly dissociated cells showing endothelial cells (black arrow), small and large steroidogenic cells (white arrows). (B) Enriched endothelial cells prior to culture (scale bar = 10 µm). (C and D) Spindle-shaped and flattened polygonal morphologies of endothelial cells in day 9 cultures (scale bar = 100 µm). (E) Lumen-like structures formed by endothelial cells in culture (scale bar = 100 µm). (F) Spontaneous formation of capillary-like structures (scale bar = 30 µm).

Figure 3 Representative images of immunostaining for BS-1 lectin (A), von Willebrand factor VIII (B), and VE-cadherin (C) in cultured luteal endothelial cells. Unstained cells (D) and cells stained with rabbit IgG (E and F) were used as negative controls. Cell nuclei were stained with DAPI (blue).

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analyzed by flow cytometry following the protocol described by Poole and Pate (2012).

LEC-induced adhesion of T lymphocytes

To validate the assay, T lymphocytes (1 × 105–1 × 106/well) were labeled with calcein AM following the manufacturer’s instructions and a standard curve was developed by reading the fluorescence on a plate reader (VictoR3; PerkinElmer) for 1 s at 485/535 nm. The correlation coefficient between the number of cells and fluorescence units (after correction for background) was 0.98. For the experiment, LEC were seeded (1 × 106 cells/well) in 96-well plates in EBM-2 medium to achieve confluence. The cells were treated with PGE2 (0.002, 0.02 or 0.2 μM), PGI2 (0.002, 0.02 or 0.2 μM), PGF2α (0.002, 0.02 or 0.2 μM) or P4 (0.5, 1, 5, 10 and 20 µM), followed by TNF (50 ng/mL) and IL1B (50 ng/mL) or PBS for the last 6 h of a 21-h incubation period. Vehicle control wells were included for each treatment. The T lymphocyte adhesion assay was conducted under static conditions. T lymphocytes (5 × 105) in 100 μL of RPMI, labeled with calcein AM (5 μM), were added to LEC

monolayers and incubated for 30 min at 37°C. Nonadhered T lymphocytes were gently removed by washing with PBS (4×). The number of T cells adhered to the LEC monolayer was determined by fluorescence (after correction for background) on the plate reader for 1 s at 485/535 nm. The data are presented in fluorescence units as a percentage of control.

Statistical analyses

Data from all experiments were analyzed by Proc mixed procedures of SAS (Statistical Analysis System Inc. version 9.3). Data generated from qPCR was subjected to analysis of covariance, using the reference gene as a covariate, or to t-test after validation that there was no effect of treatment on the reference gene. Western blot data was subjected to orthogonal polynomial regression analysis. T lymphocyte proliferation data was subjected to multiple comparisons using Bonferroni adjustment. Data generated from adhesion assays was subjected to orthogonal polynomial regression analysis and orthogonal contrasts to determine adhesion in response to increasing concentrations of hormone treatments.

Figure 4 Expression of receptors for progesterone, prostaglandins, cytokines and adhesion molecules in LEC. (A) Detection of mRNA for RPL19 and IL1B, TNF, PG and P4 receptors in midcycle bovine luteal endothelial cells ± TNF (50 ng/mL) + IL1B (50 ng/mL). Numbers 1–3 denote biological replicates of each treatment group. Mixed luteal cells (steroidogenic and endothelial; LC) were used as controls. NoRT indicates samples that were not reverse-transcribed, to control for amplification of contaminating DNA. (B) Quantification of IL1B, TNF, PG and P4 receptor mRNA in midcycle bovine luteal endothelial cells ± TNF (50 ng/mL) + IL1B (50 ng/mL); data expressed as least squares means (LSM) ± s.e.m. (n = 4). (C) Adhesion molecules (SELP and VCAM1) and CCL2 mRNA expression in early and midcycle LEC ± TNF (10 ng/mL); data expressed as LSM ± SEM (n = 4 for SELP and CCL2; n = 3 for VCAM1).

A

C

B

Day 6 Day 11–12

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In all cases, statistical significance was considered when the P value was <0.05.

Results

Characteristics of enriched LEC

The number, size, internal complexity and viability of LEC isolated by filtration and by magnetic capture were determined. The number of cells recovered using filtration was considerably greater compared to immunomagnetic separation using anti-CD31 (Fig.  1A; P < 0.05). Hence, filtration was the more efficient method of endothelial cell isolation. Analysis by flow cytometry indicated that the filtration method completely excluded cells of high internal complexity and larger size (R3; Fig. 1B). The cells in region R3 are likely luteal steroidogenic cells (LSC), while those in regions R1 and R2 are LEC. Cells recovered from the filtrate also had greater viability compared to those recovered by immunomagnetic separation (Fig.  1C; P < 0.05). To determine the relative purity of LEC isolated by filtration, lectin binding was performed on enriched LEC. Nearly 90% of enriched LEC were BS-1 lectin positive, and this proportion was significantly greater in the filtrate compared to the retentate (Fig. 1D). To further confirm the elimination of steroidogenic cells, STAR mRNA expression was determined in the retentate and the filtrate. Cells retained by the 10-µm filter tended to express greater concentrations of STAR compared to that detected in the enriched LEC fraction (Fig. 2E), but a minor proportion of steroidogenic cells remained in the filtrate.

In freshly dissociated cell preparations, LEC were smaller than presumptive steroidogenic cells, as expected, and often appeared in clusters (dark arrow, Fig.  2A). The presumptive steroidogenic cells were larger (white arrows) and exhibited an abundance of intracellular lipid (Fig.  2A), which contributed to the abundant internal complexity observed by flow cytometry (Fig. 1B). The size of enriched LEC was consistent, and about 10 µm in diameter (Fig. 2B). Enriched LEC were grown in EBM-2 medium and attachment was observed by day 3. Enriched LEC in culture exhibited both spindle- (Fig. 2C) and polygonal-shaped morphologies when cells reached >80% confluency (Fig.  2D). The two morphologies were evident following each and every isolation attempt, and often occurred within the same culture vessel. Confluent LEC, beginning on days 14–17 of continuous culture, occasionally formed lumen-like structures (Fig.  2E) and organized to form strings of connected cells characteristic of spontaneous pseudocapillaries (Fig.  2F). Cultured, enriched LEC were bound by BS-1 lectin (Fig.  3A), and expressed vWF (Fig. 3B) and VE-cadherin (Fig. 3C), consistent with characteristics indicative of endothelial cells.

Cytokine-induced expression of cytokine/hormone receptors, adhesion molecules by LEC

Luteal EC were examined for the gene expression of receptors for TNF (TNFRSF1A&B), IL1B (ILR1&2), P4 (PGR) and PG (PTGFR, PTGER2&4 and PTGIR), and the effect of the cytokines, TNF and IL1B, on receptor mRNA abundance. Although detection of some receptor

Figure 5 Adhesion molecules and cytokine mRNA and VCAM1 protein expression in luteal tissues. (A) Expression of SELP, CCL2 and VCAM1 mRNA in early and midcycle luteal tissue; data expressed as LSM ± SEM (n = 8 cows for early; n = 12 for midcycle). (B) Western blot indicating VCAM1 relative abundance in early and midcycle luteal tissue with the corresponding ACTB (n = 4). Mouse thymus tissue was used as a positive control. Numbers 1–4 denote biological replicates of each group. (C) Representative western blot of VCAM1 protein expression in luteal tissue at 0, 0.5, 1, 2, 4, 8 and 12 h after PGF2α injection with the corresponding ACTB (n = 4). (D) Quantification of western blot data for regressing luteal tissue VCAM1 protein abundance (mean ± s.e.m.; n = 4).

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mRNA was not apparent in a few individual samples of cultured cells, overall the mRNA for all receptors was detected in LEC as well as mixed luteal cells (LC; Fig. 4A). Treatment with TNF and IL1B increased the steady-state concentration of PTGIR, PTGER2 and PTGER4 mRNA, and had no effect on mRNA abundance for ILR1, ILR2, TNFRSF1A, TNFRSF1B, PTGER1, PTGFR or PGR (Fig.  4B; P < 0.05). The steady-state concentration of mRNA for the adhesion molecules, SELP and VCAM1, and the chemokine, CCL2, were quantified in LEC isolated from either early (day 6) or midcycle (days 10–12) CL. Low concentrations of mRNA were detected in untreated LEC. However TNF stimulated an increase in the abundance of CCL2 and VCAM1 (P < 0.05) and

a tendency for an increase in SELP in midcycle LEC (Fig. 4C; P = 0.087).

To determine if expression of adhesion molecules is different in developing and fully functional CL, mRNA and protein were evaluated in early and midcycle luteal tissue. Abundance of adhesion molecules and CCL2 mRNA, as well as abundance of VCAM1 protein were not different between early and midcycle luteal tissue (Fig. 5A and B). However PGF2α elicited a cubic response in VCAM1 protein abundance during the time course of luteal regression that was examined (Fig. 5C and D; P = 0.03). VCAM1 protein decreased within 30 min after PGF2α injection, followed by an increase to 4 h, then slowly declined until 12 h after PGF2α injection.

Figure 6 Effect of luteal endothelial cells (LEC) and mixed luteal cells (LC) on T lymphocyte function. (A) Representative flow cytometric plots showing the proportion of CD25+ cells in T cells cultured alone (TC), and with LEC or LC. A combination of PMA and ionomycin (PMA-Io) was used as the positive control. The gated percentage of cells for each quadrant is indicated in each plot. (B) Proliferation of T lymphocytes in response to vehicle (control), LEC and LC is shown in representative histograms. (C and D) Representative flow cytometric plots showing the proportion of CD8+ and γδ+ T lymphocytes (TC) following coculture with or without LEC. The gated percentage of cells for each quadrant is indicated in each plot. Bar graphs below each plot (A, B, C and D) indicate the quantified results of flow cytometric analysis of replicated experiments. Each experiment was repeated 3 times. Different letters indicate significant differences (P < 0.05).

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LEC-induced proliferation and phenotypic changes of T lymphocytes

The effects of LEC and mixed luteal cells (LC) on proliferation and phenotypic changes of T lymphocytes were compared. Coculture of T lymphocytes with LEC significantly increased the proportion of CD25+ T lymphocytes (indicative of activation) similar to PMA–ionomycin (positive control), but there was no effect of mixed luteal cells on CD25 expression (Fig. 6A). In contrast, mixed luteal cells induced a greater proportion of T lymphocytes to proliferate compared to LEC (Fig. 6B; P < 0.05). Finally, in assessing the effects of LEC on T lymphocyte phenotype, there was no evidence that LEC altered the proportions of T lymphocyte subsets as a consequence of coculture (Fig. 6C and D; P > 0.05).

LEC-induced T lymphocyte adhesion

There was a significant effect of P4 on T lymphocyte adhesion to LEC in both cytokine-treated and untreated LEC (Fig.  7A; P = 0.003). Low concentrations of P4 enhanced adhesion (fluorescence), while high concentrations impaired it (cubic response; P < 0.05). However the effect of prostaglandins on T lymphocyte adhesion differed between cytokine-treated and untreated cells. The highest concentrations (0.2 µM) of PGE2 and PGF2α increased adhesion of T lymphocytes

only in untreated LEC (Fig. 7B and D; P = 0.04). Similarly, 0.2 µM PGI2 tended to increase T lymphocyte adhesion only in untreated LEC (Fig. 7C; P = 0.06).

Discussion

In this study a previously untested size exclusion technique was used to isolate LEC from collagenase-dispersed CL and subsequently study the functional interactions of LEC with T lymphocytes. Filtration of dissociated, mixed luteal cells through a 10-µm filter provided greater recovery and higher viability of LEC than a previously published method utilizing magnetic cell separation. In the current study, the low yield of LEC obtained from CD31 antibody magnetic cell separation (CD31-MACS) might be attributable to the low and variable expression of CD31 among different phenotypes of LEC (Spanel-Borowski & van der Bosch 1990). Although the CD31 gene is clearly expressed in LEC of the bovine CL (Meidan  et  al. 2005), there is no previous report in which CD31 antibody has been used to isolate bovine LEC. On the other hand, anti-CD31 antibody ligated to Dynabeads has been an effective technique to isolate porcine LEC (pCL-MVECs; Zannoni et al. (2007)).

The filtration method described here takes advantage of the inherent differences in size of LEC relative to the other cell types typically found within enzymatically dispersed CL. The size of LEC from different species is estimated to be between 8 and 11 µm (Rodgers & O’Shea 1982, Lei  et  al. 1991, Fields & Fields 1996), which is less than that of small (15–23 μm) and large steroidogenic luteal cells (23–50 μm; Weber  et  al. 1987, Chegini et al. 1991, Lei et al. 1991). The purity of isolated cells was examined for the expression of BS-1 lectin-binding carbohydrates and vWF, which are specifically expressed by bovine LEC (Tuori et al. 1994, Herrman et al. 1996, Plendl et al. 1996a, Meidan et al. 2005, Cannon et al. 2007b, Grazul-Bilska et al. 2013). On the basis of vWF expression, BS1-lectin binding, size, morphology and growth characteristics, LEC isolated in this study exhibited characteristics and diversity similar to that described by Spanel-Borowski and van der Bosch (1990) and Zannoni et al. (2007). Similar diversity was not observed when endothelial cells were isolated using Ficoll (Rodgers & O’Shea 1982) or dextran separation gradients (Plendl et al. 1996a,b), nor when BS-1 lectin-linked magnetic beads were used (Meidan et al. 2005, Korzekwa  et  al. 2008). In the Spanel-Borowski and van der Bosch study (1990), the authors described isomorphic epithelioid, polymorphic epithelioid, spindle-shaped, round, and phase-dense phenotypes of bovine LEC. In the present study, not all of these shapes could be identified, but the spindle- and polygonal-shaped cells were frequently observed. According to Spanel-Borowski and van der Bosch (1990), the five

0

A B

C D

Figure 7 Effect of prostaglandins and progesterone on T lymphocyte adhesion to luteal endothelial cells. Adherence of T lymphocytes (fluorescence units) to luteal endothelial cells pretreated with (A) P4 (0–20 µM), (B) PGE2 (0.002–0.2 µM), (C) PGI2 (0.002–0.2 µM) and (D) PGF2α (0.002–0.2 µM), and treated with (black bars) or without (white bars) IL1B (50 ng/mL) and TNF (50 ng/mL). Each bar represents a mean of four biological replicates. The asterisks (*) in (A) indicate significant differences within cytokine treatment between 20 µM and both 1 and 5 µM P4 concentrations (P < 0.05) while that in (B) and (D) indicate significant differences from the control (indicated by the horizontal line). A tendency toward significance from control is indicated by + (P = 0.09).

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cell phenotypes could only be established when luteal tissue was mechanically dispersed as opposed to using collagenase digestion. Thus, the absence of certain LEC phenotypes in the current study may be due to methodology, and because growth and morphological characteristics of LEC in vitro also depend on the type(s) of growth factors present in the medium (Stolz & Jacobson 1991). Nevertheless, the total number and viability of LEC recovered after collagenase digestion and filtration in the present study is a substantial improvement from that reported by Spanel-Borowski and van der Bosch (1990). Ultimately, this technique provided sufficient numbers of cells for multiple functional studies to be performed. Additionally, because BS-1 positive cells remained detectable in the retentate, the utilization of BS-1 lectin to retrieve additional cells as well as the use of trypsin to enzymatically disperse aggregates of LEC offer further opportunities to improve yield and purity to nearly 100%.

The protein or mRNA for PTGFR (Zannoni et al. 2007, Lee et al. 2009), PTGER (Sakurai et al. 2011, Trau et al. 2015), TNFRSF1A (Okuda et al. 1999, Friedman et al. 2000, Hojo et al. 2010) and PGR (Friedman et al. 2000) has been detected in luteal endothelial cells, but the expression for some of these receptors is controversial. For instance, there are conflicting reports about the absence (Liptak  et  al. 2005) or presence (Girsh  et  al. 1996, Shirasuna et al. 2008, 2012c) of PTGFR in bovine LEC. Results from the current study tend to support the presence of transcripts for PTGFR, but the relatively low and variable abundance of these transcripts detected among LEC isolates, and the knowledge that the original filtrate of LEC remained positive for StAR expression, prevent resolving the controversy entirely. These observations notwithstanding, the overall expression of mRNA for prostaglandin and cytokine receptors, and the acute sensitivity of PGER and PTGIR mRNA to cytokine stimulation, support the concept of immune cell-LEC interaction within the CL (Walusimbi & Pate 2013), including studies that show cytokines enhance prostaglandin and chemokine effects within the CL (Nothnick & Pate 1990, Fairchild & Pate 1991, Benyo & Pate 1992, Townson & Pate 1996).

The presence of PG, P4 and cytokine receptors in LEC led us to examine if P4 and PG modulate the process of T lymphocyte adhesion to LEC, which is a prerequisite to extravasation across the endothelium. Asselin et  al. (2001) suggested that P4 promotes recruitment of eosinophils to the ovine uterus. In addition, Cid et al. (1994) demonstrated that P4 concentrations of 0.003–0.02 µM promoted adhesion of PMNs to TNF-stimulated human umbilical vascular endothelial cells (HUVECs). In contrast, P4 at 0.01 µM inhibits adhesion of leukocytes to lipopolysaccharide-treated HUVECs (Simoncini et al. 2004), and inhibits the expression of vascular cell adhesion molecule1 (VCAM1), ICAM1 (Otsuki  et  al. 2001, Piercy  et  al. 2002, Simoncini  et  al. 2004) and

E-selectin (Aziz & Wakefield 1996) in HUVECs. In the current study, the adhesion of lymphocytes to LEC pretreated with P4 was modulated relative to the concentration of P4, in a manner that was biphasic. Although the effect of P4 was independent of short-term cytokine treatment (6 h), when added after 15 h of P4 treatment, others have determined that physiological concentrations of P4 inhibit TNF-induced actions on LEC (Friedman et al. 2000). The P4-induced changes in LEC were likely through PGR (Friedman et al. 2000) and/or membrane progesterone receptors, and could have also inhibited the expression of chemokines (Goddard et al. 2013). Because the underlying mechanisms of P4 action were not determined in the current study, further studies are warranted. It would also be of interest to see if inhibition of P4 in vivo increases the adhesion and recruitment of T lymphocytes because intraluteal concentrations of P4 could be as high as 95 µM (Davis  et  al. 2010). High concentrations of P4 within the fully functional CL may limit recruitment of T lymphocytes, because more T lymphocytes, especially those that express CD8 molecules, are recruited during luteal regression when progesterone synthesis declines (Penny  et  al. 1999, Bauer  et  al. 2001, Townson  et  al. 2002, Poole & Pate 2012).

The abundance of neutrophils (de Menezes et al. 2005, Shirasuna  et  al. 2012b), eosinophils (Murdoch 1987) and T lymphocytes (Penny et al. 1999, Townson et al. 2002) in the CL is greatest during luteal regression. It is suggested that the recruitment of immune cells at this stage involves increased expression of chemokines in response to PGF2α (Haworth et al. 1998, Townson & Liptak 2003, Sales et al. 2009, Luo et al. 2011), as well as expression of adhesion molecules, such as selectin P (Shirasuna  et  al. 2012b) and intercellular adhesion molecule 1 (ICAM1; Olson et al. 2001) on endothelial cells. However immune cells are present in the functional CL (midcycle) prior to the onset of luteal regression. The role of PGE2 in recruitment of immune cells in the CL has not been studied, but PGE2 influences adhesion and transmigration of immune cells across the endothelium. For example, PGE2 enhances formylmethionyl-leucyl-phenylalanine (FMLP)-induced adhesion of leukocytes in rabbit mesenteric venules (Tromp  et  al. 2000) and increases the expression of ICAM1 (Winkler et al. 1997) on endothelial cells. In contrast, it inhibits lymphocyte binding to rat retinol and human umbilical vein endothelial cells (To & Schrieber 1990, Mesri  et  al. 1996) and transmigration across human umbilical vein endothelial cells in vitro (Oppenheimer-Marks  et  al. 1994, Mesri et al. 1996). The results of the current study support the concept of direct effects of PGF2α and PGE2 on endothelial cells to influence T lymphocyte adhesion.

As expected, TNF induced the expression of CCL2 and VCAM1, and tended to increase SELP mRNA in LEC isolated from midcycle (days 10–12) CL. Accordingly, T lymphocyte adhesion was anticipated to be greater

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in cytokine-treated than in untreated LEC. Instead we observed that T lymphocyte adhesion in response to prostaglandins was generally greater in untreated than cytokine-treated cells. This is at odds with previous studies in which cytokine-activated endothelium expressed more adhesion molecules and bound a greater number of immune cells than untreated endothelium (Carlos  et  al. 1990, Ikuta  et  al. 1991). Observing that the TNF-induced abundance of VCAM1, SELP and CCL2 transcripts occurred after a 24-h culture period, it is conceivable that the 6-h exposure to cytokines in the adhesion experiment of the current study was insufficient to translate mRNA into protein and, thus, affect adhesion of T lymphocytes to LEC. The increase in chemokine and adhesion molecule mRNA expression in response to TNF is in agreement with the results of Cavicchio et al. (2002) and Cherry et  al. (2008). The observed fluctuations in expression of VCAM1 protein in regressing luteal tissue may be due to TNF because the pattern of expression of VCAM1 during luteal regression parallels that of TNF mRNA reported previously (Petroff  et  al. 1999). Another possible explanation for the enhanced effect of prostaglandin on untreated LEC is that LEC from midcycle CL already express adhesion molecules and thus have the ability to bind T lymphocytes without additional stimulation. This possibility, however, does not resolve why the effect of the PG in vitro was less in cytokine-treated cells than untreated cells. As alluded to above, endogenous concentrations of P4 are a confounder in interpreting aspects of PG and cytokine-induced effects on adhesion of T lymphocytes. Future studies that utilize the in vitro LEC-T lymphocyte coculture model could prove insightful in this regard.

The interaction of immune cells and endothelial cells is not limited to just binding and providing a passage to tissues. During the process of extravasation, bidirectional communication between immune cells and endothelial cells occurs, resulting in T cell activation (Sancho et al. 1999). Communication between endothelial cells and T lymphocytes has mostly been studied in an allogeneic environment, in which endothelial cell-induced alloactivation induces T lymphocyte differentiation (Bedke et al. 2010, Taflin et al. 2011) and transplantation rejection (Hughes  et  al. 1990, Savage  et  al. 1993, Rollins  et  al. 1994). Recently, Spanel-Borowski (2011) suggested that cytokeratin-positive LEC may sensitize cells of adaptive immunity. The present study, in which LEC induced proliferation of autologous T lymphocytes, accompanied by increased expression of CD25, indicates activation of T lymphocytes by LEC. Activation of T lymphocytes can be induced by adhesion molecules, such as ICAM1 (Sancho  et  al. 1999), LFA3 and/or MHCII, which interact with CD2 and the T cell receptor respectively (Dustin et al. 1996, Karmann et al. 1996, Murakami et al. 1999). Although the expression of LFA3 on LEC has not been studied, bovine LEC express MHCII-DRα protein and CD80

mRNA (Cannon  et  al. 2007a,b). Expression of MHCII may be important because Cowdria ruminantium-infected bovine endothelial cells that express MHC II induce proliferation of autologous T lymphocytes (Mwangi et al. 1998). The induction of CD25 may ensure survival and response of T lymphocytes to IL2 within tissues. The lack of CD25 induction from mixed luteal cells (mainly steroidogenic cells) is paradoxical because these cells are more potent stimulators of T lymphocyte proliferation than LEC. In the proliferation assay, LEC were seeded at twice the number of mixed luteal cells, but the proliferation of T lymphocytes by luteal cells was 1.5 fold greater. Further, changes in T lymphocyte functional phenotype in response to luteal steroidogenic cells have been reported (Walusimbi & Pate 2014), but in the present study, T lymphocyte phenotype was not altered by LEC. Collectively, these results indicate that there are differences in the mechanisms by which LEC and steroidogenic cells signal to T lymphocytes, and that each of these parenchymal cell types may have a distinct role in programming of the T lymphocytes that migrate into the CL.

summary

Although endothelial cells are reported to constitute more than 50% of all cells in the CL, until now the lack of efficient methods to recover luteal endothelial cells has hindered efforts to understand the biology of these cells using cell culture. This study demonstrated that filtration is a simple, effective and efficient way of recovering large numbers of luteal endothelial cells with high viability. Initial experiments utilizing this methodology demonstrated that prostaglandins and progesterone modify adhesion of T lymphocytes to endothelial cells during extravasation and that luteal endothelial cells contribute to an environment that activates lymphocytes that take up residence in the CL.

Declaration of interest

The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.

Funding

This work was supported by National Research Initiative Competitive Grant nos. 2008-35203-04617 and 2012-67015-30212 from the USDA National Institute of Food and Agriculture.

acknowledgements

The authors would like to thank Dr D H Poole for the assistance with sample collection.

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Received 20 October 2016First decision 14 November 2016Revised manuscript received 13 January 2017Accepted 7 February 2017

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