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OPEN ACCESSJacobs Journal of Regenerative Medicine

Isolation and Characterization of Early Lineage Adult Stem Cells from the Synovial Fluid of Osteoarthritis PatientsKeith D. Crawford1, 2, *, Baldev Vasir3, Shari Benson1, Jinsoo Joo1, Kathryn A. Goldman1, Zaheed Husain4, Farnaz Hadaegh5,

Thomas S. Thornhill6

1Center for Molecular Orthopedics, Department of Orthopedic Surgery, Brigham and Women’s Hospital, Harvard Medical School,

Boston, MA, 02115, USA2Asclepius Laboratories, Inc, 27 Strathmore Road, Natick, MA, 01760, USA3Department of Medical Oncology, Dana-Farber Cancer Institute, Harvard Medical School, Boston, MA, 02215, USA4Department of Medicine, Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, MA, 02115, USA5Department of Anaesthesia, Massachussets General Hospital, Harvard Medical School, Boston, MA, 02114, USA6Department of Orthopedic Surgery, Brigham and Women’s Hospital, Harvard Medical School, Boston, MA, 02115, USA

*Corresponding author: Dr. Keith D. Crawford M.D., Ph.D., Asclepius Laboratories, 27 Strathmore Road, Natick, MA 01760, USA,

Tel: (202) 538-3336; Fax: (314) 222.6604; Email: [email protected]

Received: 03-14-2016

Accepted: 03-28-2016

Published: 03-31-2016

Copyright: © 2016 Crawford

Research Article

Cite this article: Crawford K D. Isolation and Characterization of Early lineage Adult Stem Cells from the Synovial Fluid of Osteoarthritis Patients. J J Regener Med. 2016, 2(1): 005.

Abstract

Adult stem cells (ASCs), which possess the ability to self-renew and regenerate tissue, are of significant value for the develop-ment of cellular therapies, tissue engineering tools, and drug screening models. Conventional protocols for ASC enrichment generate a small number of cells that do not represent the total ASC population of tissues. We avoided these conventional methodologies and used a different approach to identify early lineage adult (ELA) stem cells, a subpopulation of ASCs 4-6 µm in diameter, in the synovial fluid of osteoarthritic patients. These cells lack cell surface markers expressed by other ASC (e.g. CD34, CD73, CD105, SSEA-1, CXCR4). However, RT-PCR studies demonstrated expression of pluripotency genes such as NANOG, OCT4, REX1, KLF4, STELLA, and SOX. When cultured in adipogenic, chondrogenic, or osteogenic differentiation media, ELA cells differentiated into fat, cartilage, and bone tissue, respectively. Quantitative PCR analysis revealed unique molecular signatures consisting of tissue-specific and non-tissue specific mRNA in the differentiated tissues, suggesting a continuum of mRNA expression. Furthermore, the ELA cell population shared unique gene sets with embryonic stem cells, mesenchymal stem cells, and induced pluripotent stem cells. Some of these genes are unique to neuronal, cardiac, pancreat-ic, and hepatic progenitor cells, while others, such as mucins, ICAM, and tetraspans, have tissue-specific cell functions. ELA cells also demonstrated strong in vitro immunomodulatory properties by inhibiting T cell proliferation, inducing CD4+/CD25+ regulatory T cells, and inhibiting natural killer cell activity. Collectively, these observations suggest that ELA cells might be useful for cell-based regenerative therapies and the treatment of systemic diseases with immunological etiologies.

Keywords: Osteoarthritis; Synovial fluid; Bone marrow; Adult Stem Cells (ASCs); Self-Renewal; Heterogeneity; Multipo-tent, Differentiation, Transcriptome, Early Iineage Adult (ELA) Stem Cells; Molecular Signatures

Introduction

Stem cells have the remarkable capacity to self-renew, dif-ferentiate into multiple cell lineages, and reconstitute tissue in vivo [1]. Embryonic stem cells (ESCs), a pluripotent cell type, are established from early embryonic cells and possess the ability to differentiate into all three germ layers [2-5]. In contrast, adult stem cells (ASCs) are found in the developing

fetus during tissue renewal and postnatally [6,7]. Hemato-poietic stem cells (HSCs), one of the most characterized types of ASCs, have been studied for over 50 years and are known progenitors of various blood cell types [8-10]. HSCs have been used clinically to reconstitute bone marrow (BM) cells destroyed by BM ablation therapy for cancer [11,12]. There is also a heterogeneous population of non-hematopoietic stem cells in the BM. In particular, mesenchymal stem/pro-

sponse, potentially by inhibiting T cell proliferation, inducing regulatory CD4+/CD25+ T cells, and inhibiting natural killer (NK) cell activity. Thus, like MSCs, ELA cells might participate in various regenerative processes, or they might work concom-itantly with MSCs, to allow new therapies for a wide range of common and orphan diseases [23].

Materials and Methods

SF isolation and cell culture

SF was extracted from the knees of patients diagnosed with os-teoarthritis (OA) following appropriate institutional approved protocols. Within 24 h of harvest, the SF was diluted 10:1 in di-lution buffer (phosphate-buffered saline (PBS) supplemented with 10% fetal bovine serum (FBS; HyClone, Logan, UT)) and 10 mM ethylenediaminetetraacetic acid (EDTA; Gibco, Grand Island, NY). To extract the cellular component, the diluted SF was spun at 500 x g for 30 min, and the pellet was resuspend-ed in dilution buffer. This process was repeated twice at 300 x g for 30 min, and the final pellet was resuspended in Hank’s balanced salt solution (HBSS; Gibco). The pelleted cells were either directly analyzed or subjected to culture expansion and differentiation. Samples that underwent culture expansion and differentiation were suspended in growth medium (MSCGM™ Human Mesenchymal Stem Cell Growth BulletKit™ Medium or MSCGM-CD™ Mesenchymal Stem Cell Chemically Defined Me-dium with or without 1% FBS; Lonza, Basel, Switzerland) and plated at a concentration of 3000 cells per cm2 (225,000 cells total) in a T-75 vented tissue culture flask (BD Biosciences, San Jose, CA). All culture media were supplemented with 100 U/ml penicillin and 1000 U/ml streptomycin (PCN-Strep; Gibco) and exchanged every 48 h. Once cells reached >80% conflu-ence, they were harvested with Trypsin-EDTA and replated into new T-75 flasks. Cells were cultured at 37oC with 5% CO2for all experiments. Samples were either immediately seeded into cell culture or mixed 1:1 with freezing medium composed of Dulbecco’s Modified Eagle’s Medium (Gibco) supplemented with 20% FBS and 20% dimethylsulfoxide (Sigma-Aldrich) and stored in liquid nitrogen at less than -150°C.

Flow cytometric characterization of ELA cells

Antibody use was based on the minimal surface marker pan-el proposed by the International Society of Cellular Therapy [16]. Fluorochrome-labeled antibodies against the following markers and matching isotype controls were obtained from BD Biosciences: CD44-PE, CD45-PE, CD49-PE, CD105-PE, CD133-PE, CD34-PE (clone 581), CD73-PE, CD90-PE, CD99-PE, CD235a-PE, MUC1-PE, HLA Class1-PE, HLA-DR-PE, SSEA1-PE, HLA-DR-PE, IgG1-PE, IgG1-FITC, IgG2a-PE, IgG2bkappa-PE, IgM-PE, CD4-FITC, CD8-FITC, CD69-PE, CD3-FITC, and CD25-PE. Anti-CXCR4-PE (CD184) antibody was obtained from R&D

genitor cells (MSCs) are also thought to originate from the BM and comprise 0.01-0.001% of nucleated BM cells [13]. MSCs are found in the peripheral blood, umbilical cord blood, adi-pose tissue, skeletal muscle, liver, lungs, synovium, dental pulp, apical papilla, amniotic fluid, and fetal blood [14]. Because MSCs are found in extremely low numbers in the BM, sustained ex vivo culture on tissue culture plastic is required to generate sufficient cell numbers for phenotypic characterization [15]. MSCs most commonly express surface markers such as CD29, CD44, CD49a-f, CD51, CD73, CD105, CD106, CD166, and Stro1 and lack expression of hematopoietic lineage markers such as CD11b, CD14, and CD45 [16]. MSCs are multipotent ASCs capa-ble of differentiating into various mesodermal tissues, such as adipose, cartilage, and bone [16]. Other groups have reported that MSCs are capable of differentiating into ectodermal and endodermal tissues, such as lung, skin, pancreas, and liver tis-sue [17]. It has been hypothesized that MSCs may not directly repair tissue but instead secrete trophic factors that decrease cell death, recruit immune cells to the injury site, and promote healing [18,19]. Presently, MSC immunomodulatory properties are being assessed in clinical trials to determine their efficacy in treating a variety of immune-related diseases [20].

Most ASC studies focus on BM-derived stem cells and use discontinuous density gradients, such as Ficoll-Paque and Lymphoprep, and plastic adherence to enrich for ASCs [21]. Although density gradients effectively separate debris, plate-lets, and red blood cells (RBCs) from the mononuclear cells in the buffy layer, they also inadvertently discard a subset of ASCs [22]. To avoid potential discrepancies in the isolation of ASCs, we decided to forgo the use of discontinuous gradients and prolonged culture on tissue culture plastic to harvest ASCs. We chose synovial fluid (SF) as a source due to low RBC contami-nation. To isolate ASCs, we used time sedimentation of diluted SF. Cells in the enriched ASC population measured 4-6μm in diameter (mean = 5μm). Flow cytometry and gene expression analysis suggested that this ASC population expresses genes and proteins generally thought to be restricted to ESCs. In ad-dition, these cells did not express MHC class II, CD44, CD45 or CD49, but unlike our prior studies they were smaller and had minimal MHC class I expression. Semi-quantitative PCR studies showed expression of embryonic transcription factors such as OCT4, REX1, NANOG and SOX2, suggesting pluripoten-cy. We therefore named these cells early lineage adult (ELA) stem cells, since they shared the gene expression profiles of the cells described in earlier studies.

In this study, we investigated the ability of ELA cells to self-re-new and differentiate into multiple lineages. Moreover, we optimized the isolation, culture, and expansion conditions for these cells in vitro. We show that ELA cells can differentiate into adipose, cartilage, and bone lineages, and that they also express genes from other cell types. In addition, we deter-mined that ELA cells are potent modulators of the immune re-

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Systems (Minneapolis, MN). Anti-PD1-PE (CD279) antibody was obtained from eBiosciences (San Diego, CA). Upon con-fluence, cells from one 75 cm2 flask were harvested, washed, and counted. Cells were kept on ice and suspended in incuba-tion buffer (Dibco’s PBS + 2% FBS + 1 mM EDTA). After cen-trifugation and aspiration of supernatant, 5 or 10 μl of anti-body (depending on cell number) was applied directly onto the pellet. Cells were incubated at 4°C for 30-45 min, washed, resuspended, and analyzed in a FACSCalibur machine using CellQuest™ software (BD Biosciences). The pluripotent prop-erties and ESC marker status of ELA cells were determined by intracellular staining using monoclonal antibodies against OC-TA4-PE, RUNX2-PE, SOX9-PE, REX1-PE, NANOG-PE, and KLF4-PE with matching isotype controls and by RT-PCR of freshly isolated and culture-expanded cells. For intracellular staining, cells were permeabilized with Cytofix/Cytoperm solution (BD Biosciences) and thereafter subjected to intracellular staining with the appropriate antibodies.

Self-renewal properties of ELA cells

ELA cell cultures grew as monolayers. Following cell sorting, both marker-positive and marker negative cultures were set up in parallel. At days 3 and 5, cultures were rinsed with PBS, detached with trypsin-EDTA, centrifuged, and resuspended in media. Duplicate aliquots were placed into 96-well plates, and 10 μl of Cell Counting Kit-8 solution (Dojindo Molecular Technologies Inc., Gaithersburg, MD) was added to each well. Following 3 h incubation at 37°C, A450 was measured using a Victor5 Light Luminescence Counter (PerkinElmer Life Sci-ences, Boston, MA) and compared with standards of known cell numbers. To detect apoptotic cells, cultures were fixed and stained with the fluorescence-based ApoAlert DNA Fragmen-tation Assay Kit (BD Biosciences) following the manufacturer’s protocol.

Adipogenic, chondrogenic, and osteogenic differentiation of cells isolated from synovial fluid

ELA cells were suspended in chemically defined media with 1% FBS, passaged upon reaching 80% confluence, and plat-ed in 75cm2 vented cell culture flasks at a concentration of 150,000/cm2, with a total volume of 25 ml per flask. After 20 passages, cells were plated into 12-well plates at a concentra-tion of 200,000 cells/well and cultured in the appropriate dif-ferentiation media. For adipogenic differentiation, cells were cultured in StemPro Adipocyte Differentiation Media (Invit-rogen) supplemented with PCN-Strep at a total volume of 1.5 ml/well. The media was changed every 48 h. After 21 d, the cells were harvested for histochemical staining and real-time quantitative PCR (qPCR). Differentiated cells were stained with fresh Oil Red O solution (Sigma-Aldrich) to verify adi-pocyte characteristics. For chondrogenic differentiation, cells were cultured in Osteocyte/Chondrocyte Differentiation Basal

Medium (Invitrogen) with Chondrogenesis Supplement (Invit-rogen). These cell cultures were then stained with Alcian Blue (Sigma-Aldrich) for chondrocyte detection. For osteogenic differentiation, cells were cultured in Osteocyte/Chondrocyte Differentiation Basal Medium (Invitrogen) with Osteogenesis Supplement (Invitrogen). To assess the presence of osteo-blasts, cell cultures were stained with 5-bromo-4-chloro-3-indolyl-phosphate/nitro blue tetrazolium (NBT/BCIP; Invit-rogen). In all three differentiation studies, positive cells were assayed by counting 50-100 cells in multiple fields using light phase microscopy.

RNA isolation and RT-PCR

Total RNA was extracted from freshly isolated and culture-ex-panded undifferentiated cells as well as differentiated cells. Total RNA was purified using TRIzol® reagent according to the manufacturer’s protocol (Invitrogen). The same source of RNA was used for RT-PCR and DNA microarray analysis (see below). First-strand cDNA was obtained by reverse transcrip-tion using 3 mg total RNA according to the manufacturer’s in-structions (Invitrogen). Primer sequences are shown in Table 1. PCR products were electrophoresed on 1.5% agarose gels to verify DNA fragment sizes. For DNA microarray analysis, cDNA was synthesized using the SuperScriptTM III First-Strand Syn-thesis System (Miltenyi Biotec, San Diego, CA). RT-PCR assays were performed using qPCR Mastermix Plus for SYBR Green (Miltenyi Biotec) according to the manufacturer’s protocol (see below). For normalization, differential levels of gene expres-sion were calculated in relation to beta actin and expressed as a ∆CT value, as previously described [24].

Preparation for DNA microarray analysis

Total RNA was extracted from synovial ELA cells in a mono-layer of human synovial ELA cells cultured for 3 d. Cells were lysed using the SuperAmp preparation kit and delivered to Miltenyi Biotec on dry ice. SuperAmp RNA amplification was performed according to Miltenyi Biotec’s protocol based on a global PCR protocol. mRNA was isolated using magnetic bead technology. Amplified cDNA samples were quantified using an ND-1000 Spectrophotometer (NanoDrop Technologies), and 250 ng of each cDNA were used as template for Cy3 and Cy5 labeling according to Miltenyi Biotec’s protocol. Cy3- and Cy5-labeled cDNAs were combined and hybridized for 17 h at 65°C to the Agilent Whole Human Genome Oligo Microarray 4x44K probe set using Agilent’s recommended hybridization chamber and oven. Control samples were labeled with Cy3 and experimental samples were labeled with Cy5.

Data processing and analysis

Feature Extraction Software (Agilent) was used to read and

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ELA cells were co-cultured with T cells labeled with 5-6-car-boxyfluorescein diacetate succinidyl ester (CFSE; Cell Trace Cell Proliferation Kit; Molecular Probes/Invitrogen Life Tech-nologies) in 96-well plates at 1:10, 1:20, and 1:40 ratios in trip-licate, along with non-ELA cell controls. T cell proliferation was stimulated with CD3/CD28 and analyzed with flow cytometry for CFSE fluorescence after 5 d. Immunosuppressive proper-ties of MSCs (Lonza, Walkerville, MD) were assayed in parallel with the same methods. Flow cytometry data was analyzed us-ing FlowJo software (Ashland, OR) to obtain the Proliferation Index (PI). T cell suppression for each sample was calculated as (1- ([PI with ELA cells]/[PI without ELA cells]) x 100. As a final assay for immunosuppression, freshly isolated ELA cells and PBMCs were co-cultured 1:10 in 96-well plates for 5-7 days, harvested into 5 ml tubes, and labeled with a combina-tion of directly conjugated antibodies as follows: CD4-FITC/CD25-PE; CD8-FITC/CD25-PE; CD3-FITC/CD69-PE, and CD3-FITC/PD1-PE, as well as matching isotype controls. The per-centages of CD4+ or CD8+ T cells expressing CD25, and CD3+ T cells expressing CD69 or PD1 were determined by bi-dimen-sional FACS analysis.

ELA cell suppression of NK cell activity was determined by a chromium release assay [29]. NK cells were co-cultured with equal numbers of ELA cells in RPMI 1640 tissue culture media (Mediatech, Herndon, VA) supplemented with 10% pooled hu-man AB serum, antibiotics, and cytokines for 24 h. Following incubation, NK cells were transferred to wells for co-culture with chromium-labeled K562 target myeloma cells (ATCC, Manassas, VA) at 10:1, 5:1, 2.5:1, and 1:1 ratios. Specific cyto-toxicity was calculated as 100 x (experimental-spontaneous)/ (maximum-spontaneous).

Statistical analysis

Results are expressed as mean ± SEM. Statistical comparisons were performed using the Student’s t-test. P values <0.05 were considered statistically significant.

Results

Cellular phenotypes of SF cells

SF contains a wide variety of mononuclear cells, some of which revealed a forward and side scatter flow cytometry pattern similar to peripheral blood. This pattern consists of four re-gions, three of which represent neutrophils, myeloid cells, and lymphocytes (Figure 1A). The fourth region, much smaller in size and side scatter, was previously thought to primarily rep-resent cell debris and RBCs (Figure 1B). We found that this population had less fluorescence compared with other regions and <200 forward scatter (FSC) linear units (Figure 1A). Anal-ysis of this cell population from 5 OA patients revealed a mean

process the microarray image files and raw datasets. These datasets, together with publically available datasets from the NIH Gene Expression Omnibus (GEO), were exported to JMP software (SAS Institute Inc., Cary, NC) for further analyses. The input datasets were transformed into log base 2, and row-by-row statistics were computed. Datasets were normalized to the median global intensity [25,26].

Hierarchical clustering and functional analysis

To identify genes expressed at high levels in ELA cells, an un-supervised hierarchical clustering was performed on the nor-malized dataset. JMP Software was used to perform and visu-alize this clustering. One-way ANOVA was performed on the data obtained from the hierarchical clustering, and a volcano plot was generated to represent the intensity ratio for each gene in ELA cells and MSCs. The x-axis displays the log2 ratio of the gene intensities. A log2 ratio of 1corresponds to approx-imately two-fold change. The y-axis shows the –log10 (p-value) for the comparison between ELA cells and MSCs. Genes that were differentially expressed in ELA cells and MSCs were iden-tified. These genes, along with their fold-change values, served as the input to the Ingenuity Pathway Analysis (IPA®, Qiagen) program. Differently expressed genes were uploaded into the IPA application and used as the starting point for generat-ing biological networks [27]. A right-tailed Fisher’s test with α=0.05 and the whole database as a reference set was used to determine if the enrichment of genes with particular biological functions or molecular processes was significant.

Immunomodulatory properties of ELA cells

The immunosuppressive properties of ELA cells were assayed by several methods. Suppression of T cell proliferation was determined by in vitro co-culture experiments performed in triplicate. ELA cells were irradiated in suspension with a dose of 30 Gy prior to coculture using the Gamma cell Elan device (Best Theratronic, Ottawa, ON, Canada). Irradiated and non-ir-radiated ELA cells, either freshly isolated or cryopreserved and then cultured for 24-48h, were co-cultured with freshly isolat-ed human peripheral blood mononuclear cells (PBMCs) at a 1:10 ratio for 5 d at 37°C. ELA cell suppressive function was determined by a [3H]-Thymidine (1μCi/well; 37kBq; NEN-Du-Pont) uptake assay, as previously described [28]. Data are ex-pressed as counts per minute (cpm) or as a stimulation index (SI). SI was determined by calculating the ratio of experimen-tal [3H]-Thymidine incorporation to background [3H]-Thymi-dine incorporation by unstimulated T cells. These experiments also assessed if allo-reactive T cells were stimulated by ELA cells, which would be indicated by higher counts in a prolifer-ation assay.

To further demonstrate their immunosuppressive properties,

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viability of 94 ± 0.65% and a mean cell size of 5.9 ± 0.31 μm(range: 4-8 μm) (Figure 1C).

Figure 1. Identification of early lineage adult (ELA) stem cells in the synovial fluid (SF) of patients with osteoarthritis (OA). (A) Representative forward- and side-scatter profiles of mononuclear cells isolated from the SF of an OA patient indicating the location of a small cell population in relation to other cell types. (B) Forward- and side-scatter profile of a gated small population of cells depicting a het-erogeneous population with a varied cell size and scatter profile. (C) Viability and cell-size determination of a gated small cell population using the Roche CASY Cell Counter and Analyzer System. (D) Expres-sion of pluripotent intracellular and surface markers, as determined by FACS analysis of a gated population of small cells in Group 1, Group 2, and Group 3. (E) RT-PCR analysis of pluripotency marker expres-sion in a small cell population isolated from three separate samples of SF, with NTERA-2 cells (a stem cell line) as a positive control. Prim-ers are specific for transcripts from the respective endogenous locus. GAPDH was used as a loading and internal control.

We next evaluated the phenotypic characteristics of these cells by staining for proteins associated with peripheral blood mononuclear cells and ASCs. Further flow cytometric analysis showed three distinct subgroups of cells (Figure 1B). CD45 and CD235a, surface markers of leukocytes and RBCs, respec-tively, were absent from this cell population. MHC Class I, a protein found on all cells except RBCs and immature stem cells, was observed on a subset of this group (Figure 1D). Further-more, we observed very high expression of MUC1 and CD99 on this cell population. Further analysis revealed no expres-sion of CD73, CD90, CD105, CD133, CXCR4, or SSEA-1 on any of the subgroups. However, intracellular staining with directly conjugated monoclonal antibodies revealed high expression of REX1 and varying degrees of OCT4, NANOG, SOX9, and RUNX2 expression (Figure 1D). RT-PCR analysis from three patients revealed that most of the tested pluripotency-associated mR-

NAs were expressed in this small cell population, with REX1 being the most highly expressed (Table 1, Figure 1E). Notably, we identified a splicing variant of NANOG, which might suggest a greater diversity of self renewal and pluripotency proteins. The Ntera cell line was used as a positive control in these stud-ies [30], and a sample lacking reverse transcripts was used as a negative control.

ELA cell growth and self-renewal

In order to investigate and optimize cell growth, ELA cells were cultured in three different media types; standard expansion medium, chemically defined medium (CD), and CD supple-mented with 1% FBS. ELA cells cultured in standard medium reached 90% confluence in 8 days for donor A and 7 days for donor B, generating 2.8 x 106 and 2.2 X 106 cells, respectively. Doubling times for both samples were greater than 120 h. In contrast, when ELA cells were cultured in CD in the absence or presence of 1% FBS, higher yields were generated with short-er doubling times. Notably, CD supplemented with 1% FBS appeared to be the optimal media for self-renewal capacity, generating yields of 31.5 x 106 (donor A) and 21.2 x 106 (donor B) ELA cells with doubling times of 19h and 27h, respectively (Figure 2A-B). We therefore chose this culture media for all future experiments. In CD media with 1% FBS, cell morpholo-gy was round immediately after plating but became elongated and spindle-like within 4 d (Figure. 2C-D).

Figure 2. Cell culture and self-renewal properties of early lineage

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Table 1 Primer sequences for RT-‐PCR amplifica8on of target genes

Gene Primer Gene Bank accession number Product size (bp)

GAPDH F: 5'-‐ AGCCACATCGCTCAGACAC-‐3' NM_001256799.1 66

R: 5' -‐ GCCCAATACGACCAAATCC-‐3'

NANOG F: 5' -‐ TGTCTTCTGCTGAGATGCCT-‐3' NM_024865.2 88

R: 5' -‐ TCTCTGCAGAAGTGGGTTGT-‐3'

SOX2 F -‐ 5' -‐ AGCTCGCAGACCTACATGAA-‐3' NM_003106.3 151

R: 5' -‐ TGGAGTGGGAGGAAGAGGTA-‐3'

OCT4. F: 5' -‐ ACATGTGTAAGCTGCGGC C-‐3' NM_002701.4 297

R: 5' -‐ GTTGTGCATAGTCGCTGCTTG-‐3'

REX1 F: 5' -‐ GGATCTCCCACCTTTCCAAG-‐3' NM_020695.3 105

R: 5' -‐ GCAGGTAGCACACCTCCTG-‐3'

GDF3 F: 5' -‐ TGCTGTTCACTTCAACCTGC-‐3' NM_020634.1 156

R: 5' -‐ AGGGAGCATCTTAGTCTGGC-‐3'

STELLA F: 5' -‐ GGAAGCTTTACTCCGTCGAG-‐3' NM_199286.3 (AY230136.1) 65

R: 5' -‐ GCCACTCATCTTCGATTTCC-‐3'

KLF4 F: 5' -‐ CGTTGACTTTGGGGTTCAGG-‐3' NM_004235.4 139

R: 5' -‐ GCGAACGTGGAGAAAGATGG-‐3'

adult (ELA) cells in vitro. (A) Bar graph depicting growth of ELA cells from two separate donors cultured in standard culture media or chemically defined (CD) culture media with or without 1% fetal bo-vine serum (FBS). (B) Phase micrographs demonstrating pattern and density of ELA cell growth at two different magnifications. (C) Phase micrographs exhibiting the pattern and density of ELA cell growth at days 1, 4, and 7 in CD media supplemented with 1% FBS from three separate donors. (D) Total ELA cell counts from three separate do-nors cultured in CD media with 1% FBS at different passage numbers. Numbers in parenthesis represent population doubling time during labeled passage growth period.

ELA cell differentiation

The differentiation potential of ELA cells was investigated by culturing cells under conditions that favored adipogenic, chondrogenic, and osteogenic differentiation. Cells cultured in Adipocyte Differentiation Media for 21 d formed vacuoles that stained positive for Oil Red O, a fat-soluble dye that stains lipids (Figure. 3A). Control cells showed no incorporation of Oil Red O. Cells cultured for 21 d in Chondrocyte Differentia-tion Media exhibited diffuse Alcian Blue staining, suggesting production of acid mucopolysaccharides and glycosaminogly-cans normally found in cartilage (Figure 3B). Cells cultured in Osteocyte Differentiation Media and stained with NBT/BCIP revealed flat, purple cell bodies (Figure 3C). NBT/BCIP is con-verted into purple stain by alkaline phosphatase, an enzyme found in osteoblasts. No enzymatic activity was observed in control cells.

Figure 3. Differentiation of early lineage adult (ELA) cells to ad-ipocytes, chondrocytes, and osteocytes. The differentiation po-tential of ELA cells was investigated by culturing cells for 21 d under conditions that favored adipogenic, chondrogenic, or osteogenic dif-ferentiation. (A) Adipogenic differentiation was indicated by accumu-lation of neutral lipid vacuoles that stained with Oil Red O. (B) Chon-

drogenic differentiation was assayed with Alcain Blue, which labels the acid mucopolysaccharides and glycosaminoglycans of cartilage. Diffuse blue staining was observed throughout the slide. (C) Osteo-genic differentiation was assayed with BCIP/NBT, a substrate that turns purple in the presence of alkaline phosphatase. Uninduced cells were used as negative controls in the differentiation experiments. Total RNA was extracted from these differentiated cells, and cDNA derived from mRNA was amplified based on the global PCR protocol described in Materials and Methods. (D-F) Real time RT-PCR analysis of selected specific genes expressed in differentiated ELA cells. The expression of genes was compared to the expression of beta-actin as an internal control and the values expressed as ΔCT. Negative bars in-dicate a decrease in expression of that particular gene. (D) ELA cells differentiated into adipocytes. (E) ELA cells differentiated into chon-drocytes. (F) ELA cells differentiated into osteocytes.

In order to further investigate the extent of induced ELA cell differentiation, cells were harvested to investigate the pres-ence of adipogenic, chondrogenic, and osteogenic genes by qPCR. Cells from the adipogenesis conditions showed high expression of the adipocyte lineage genes PPARG-tv1, PPARG-tv2, LPL, FABP4, ADIPOQ, LEP, PLIN, and CFD (Figure 3D). Ad-ipogenesis-specific genes were also detected in chondrogenic and osteogenic conditions. Cells from the chondrogenic con-ditions showed high expression of the chondrocyte lineage genes BGN, DCN-tvA2, ANXA6-tv2, MMP13, SRY, and COMP and low/absent expression of MATN1 and COL2A1 (Figure. 3E). These genes were also detected at similar levels in cells undergoing adipogenesis or osteogenesis. Cells from the os-teogenic conditions showed high expression of the osteocyte lineage genes RUNX2-tv3, RUNX2-tv1, RUNX2-tv2, and PHEX, similar to the chondrogenic conditions. RUNX2.2 and PHEX were also detected in adipogenesis conditions. Low/absent expression of BGLAP, SPP1, SPP2, and SPP3 was also observed in the osteogenic conditions (Figure 3F). There was low/ab-sent expression of all lineage-specific genes in control cells. Although cells were cultured in specialized media for differ-entiation, the differentiation process was not absolute because cells also expressed genes from other lineages (Figure 3D-F). These observations suggest that detailed studies are necessary to determine optimum culture conditions for stem cell differ-entiation, and that molecular signatures may be necessary to define the differentiation process.

Large-scale gene expression profiling of freshly isolated, un-differentiated ELA cells showed expression of differentially expressed progenitor and tissue-specific genes with diverse functions (Table 2). These profiling studies suggested the po-tential of ELA cells to differentiate into other lineages, such as neuronal, cardiac, pancreatic, and liver cells (Tables 2 and 3). Moreover, expression of genes with specific cellular functions, such as mucins, ICAM, tetraspans, and collagens (Table 3), sug-gests that ELA cell phenotype may not be tissue specific. In-stead, ELA cells might be a heterogeneous population of multi-potent cells with the capacity to differentiate into endodermal, mesodermal, and ectodermal cells.

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Freshly harvested ELA cells (small population) was isolated from SF of OA patient. Total mRNA from the sample was used for microarray analysis.

Table 2. Selected panel of differentially expressed progenitor-‐ and tissue-‐speciDic genes from freshly-‐isolated ELA cell population

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Gene Gene Description Chr. Location Protein Expression

COL4A1 Collagen, type IV, alpha 1 13q34 Secreted Positive expression in endothelium and in basal membranes of most epithelial

tissues

COL4A2 Collagen, type IV, alpha 2 13q34 Secreted Positive expression in alveolar cells, muscle and stromal cells including, in addition

to the small intestine, gall bladder and parts of pancreas.

COL8A1 Collagen, type VIII, alpha 1 3q12.1 Secreted High level of expression in internal elastic lamina of blood vessels and cytoplasmic

and membranous locations premenopausal uterine glands. Connectie tissue,bronchus, renal glomeruli, smooth and skeletal muscles and Langerhans cells also

stain positive

COL9A2Collagen, type IX, alpha 2

1q34.2 Secreted High expression in gastrointestinal tract, renal tubules and chondrocytes and low

expression hepatocytes.

COL9A3 Collagen, type IX, alpha 3 20q13.33 Secreted Most normal tissues show positive cytoplasmic staining. Glomerular whiletrophoblastic, and glial cells stain weakly or are negative.

COL13A1 Collagen, type XIII, alpha 1 10q22.1 Secreted Expressed in breast, trophoblastic cells, salivary gland, seminal vesicle, urothelialcells, pancreatic ducts and squamous epithelial tissue Fallopian tube, gall bladder

and bile duct cells were moderate positivity.

COL14A1 Collagen, type XIV, alpha 1 8q24.12 Secreted Squamous and respiratory epithelia, hepatocytes, urinary and gall bladder,

endometrium, Leydig cells gastrointestinal tract and prostate showed moderate.

COL15A1 Collagen, type XV, alpha 1 9q22.33 Secreted Expressed predominantly in internal organs such as adrenal gland, pancreas and

kidney and localized to basement membrane zones, functioning to adhere basement

membranes to underlying connective tissue stroma.

COL27A1 Collagen, type XXVII, alpha 1 9q32 Secreted Involved in the calcification of cartilage and the transition of cartilage to bone

MUC1 Mucin 1, 1q21 Cell Surface

Associated

Expressed on the apical surface of epithelial cells, especially of airway passages,

breast and uterus.

MUC2 Mucin 2 11p.15 Secreted Expressed in colon, small intestine, colonic tumors, bronchus, cervix and gall bladder

.

MUC3A Mucin 3A 7q22.1 Cell Surface

Associated

Expressed in small intestine, colon, colonic tumors, heart, liver, thymus, prostate,

pancreas and gall bladder.

MUC4 Mucin 4 3q29 Cell Surface

Associated

Expressed in the thymus, thyroid, lung, trachea, esophagus, stomach, small intestine,

colon, testis, prostate, ovary, uterus, placenta, and mammary and salivary glands.

MUC5AC Mucin 5AC 11p15.5 Oligomeric

Mucus/Gel‐Forming

Secreted by gastric and respiratory tract epithelia, which protects the mucosa from

infection and chemical damage.

MUC6 Mucin 6 11p15.5 OligomericMucus/Gel‐

Forming

Expressed in the regenerative zone of gastric antrum, gastric body mucosa andgastric incisura mucosa.

MUC7 Mucin 7 4q13.3 Oligomeric

Mucus/Gel‐

Forming

Expressed in mucous acinar cells of salivary gland tissues and thought to play a role

in removal of bacteria in the oral cavity aid in mastication, speech, and swallowing.

MUC8 Mucin 8 12q24.33 Cell Surface

Associated

Expressed in the human airway mucin

MUC13 Mucin 13 3q21.2 Cell Surface

Associated

Expressed in gastrointestinal and respiratory tracts

MUC15 Mucin 15 11p14.2 Cell Surface

Associated

Spleen, thymus, prostate, testis, ovary, small intestine, colon, peripheral blood

leukocyte, bone marrow, lymph node and lung

MUC17 Mucin 17 7q22.1 Oligomeric

Mucus/Gel‐Forming

Expressed almost exclusively in the intestine.

Encodes a protein that functions as a cytoprotectant, maintains luminal structure,and provide signal transduction

MUC19 Mucin 19 12q12 OligomericMucus/Gel‐

Forming

Expressed by corneal epithelial cells, mucous cells of the submandibular gland andsubmucosal gland and trachea middle ear epithelial cells.

MUC20 Mucin 20, Cell Surface

Associated

3q29 Expressed kidney, placenta, lung, prostate, liver, and digestive system.

ICAM1 Intercellular Adhesion

Molecule 1 (CD54)

19p13.2 Cell Surface

Associated

Interacts with ntegrins of type CD11a / CD18, or CD11b / CD18


Molecule 2 (CD102)

17q23.3 Cell Surface

Associated



Molecule 3 (CD50)


Associated



Molecule 4


Associated

Landsteiner‐Wiener (LW) blood group antigen(s) and that shares similarity with the

intercellular adhesion molecule (ICAM) protein family


Molecule 5, Telencephalin


Associated

Expressed in Cerebral cortex, cerebellum, hippocampus, and lateral ventricle, and on

the surface of telcephalic neurons and key for neuronal development

TSPAN4 Tetraspanin 4 11p15.5 Cell Surface

Associated

Expressed in multiple tissues but is absent in brain, lymphoid cells, and platelets.

TSPAN6 Tetraspanin 6 Xq22 Cell Surface

Associated

Cytoplasmic and membranous staining was observed in most normal tissues. .

TSPAN10 Tetraspanin 10 17q25.3 Cell Surface

Associated

Expressed in the eye including iris, ciliary body, retinal pigment epithelium, but not

lens


Associated

Expressed in the adrenal gland, Leydig cells, bone marrow, heart, spleen, neurons,

skeletal muscle


Associated

Expressed in gall bladder, prostate and glandular cells

TSPAN16 Tetraspanin 16 19p13.2 Cell Surface

Associated

Expressed in pancreas, bronchus, gall bladder, renal tubules, fallopian tube, Leydig

cells, hematopoietic cells and myocytes.


Associated

Expressed in pancreas, liver, breast, prostate, and neuronal cells

UPK1A

(TSPAN21)

Uroplakin 1A 19q13.12 Cell Surface

Associated

Expressed peripheral nerves, umbrella cells in urothelium, thyroid gland, Purkinje

cells, stomach, and adrenal glandular cells.

ROM1 Tetraspanin 23 11q12.3 Cell Surface

Associated

Expressed in the photo receptor disk rim of eye

CD151 Tetraspanin 24

(Rod outer segmentmembrane protein)


Associated

Expressed in blood vessels, and epidermis, but essential for proper assembly of the

glomerular and tubular basement membrane of the kidney

Table 3. Expression of selected group of genes for secreted proteins (collagens/mucins) and surface-‐expressed proteins (mucins, ICAMS, and tetraspans) in freshly-‐isolated ELA cells

ELA cells were isolated from SF of OA paEents and the total RNA was extracted for microarrary analysis. (Chr. Chromosome)

DNA microarray analysis

To compare ELA cells and MSCs, we examined their respec-tive gene expression profiles in triplicate by microarray. The correlation coefficient between these microarray datasets ob-tained from repeated experiments was greater than 0.98, in-dicating high reproducibility. The entire set of expressed pro-tein-coding genes was used for a non-supervised hierarchical clustering analysis. The dendrogram in Figure 4A shows that freshly isolated and frozen/expanded ELA cells isolated from the same tissue were found in different clusters, whereas the three categories of MSCs (BM-derived, CD105+, and CD133+) clustered together. These results suggested that ELA cells have a gene expression profile distinct from MSCs.

Figure 4. Comparison of gene expression profiles of early lin-eage adult (ELA) cells and mesenchymal stem/progenitor cells (MSCs). Hierarchical cluster analysis of qRT-PCR data was performed on freshly isolated and cultured/expanded ELA cells and bone mar-row (BM)-derived, CD105+, and CD133+ MSCs. Expression levels were normalized to β-actin. Black represents 1, red represents >1, green represents <1, and grey represents below detection limits. (A) Dendrogram comparing gene expression in primary ELA cells, BM-derived MSCs, CD105+ MSCs, and CD133+ MSCs. (B) Dendrogram of a hierarchical cluster analysis comparing genes expressed in ex-panded ELA cells against MSC gene data sets available from the NIH gene Expression Omnibus in addition to those generated in this study. (C) Volcano plots comparing specific genes upregulated in primary ELA cells and expanded ELA cells (upper left), BM-derived MSCs (up-per right), CD105+ MSCs (bottom left), and CD133+ MSCs. Genes ex-pressed above the broken red line represent genes specific to primary ELA cells on the left and genes specific to MSCs on the right. (D) Venn diagram comparing upregulated genes in expanded ELA cells, BM-de-rived MSCs, CD105+ MSCs, and CD133+ MSCs. (E) Venn diagram de-picting the percentage of genes expressed in expanded ELA cells that

overlap with other categories of stem cells, utilizing published gene datasets available from the NIH Gene Expression Omnibus.

To verify that ELA cells are a distinct population of cells com-pared with MSCs, we utilized high-density oligonucleotide microarrays and functional network analysis. DNA microar-ray analysis was used to identify genes expressed in ELA cells, and the results were compared with datasets from the NIH GEO and our laboratory (Figure 4B). This analysis revealed that 25% of the genes expressed by ELA cells were shared by BM-derived MSCs (Figure 4B). The results were visually repre-sented by a volcano plot to compare specific genes upregulated in ELA cells and MSCs (Figure. 4C). Additionally, we generated Venn diagrams to compare gene expression in ELA cells and BM-derived, CD105+, and CD133+ MSCs (Figure 4D). From this analysis, we concluded that ELA cells are a distinct pop-ulation of ASCs.

DNA microarray analysis identified genes specific to MSCs, ESCs, and induced pluripotent stem cells (iPSCs) in the ELA cell population (Figure 4E). For example, ESCs expressed 1460 genes that were not expressed by the other stem cell types. ELA cells expressed unique genes as well as genes in common with ESCs, MSCs, and iPSCs (Table 4). Furthermore, ELA cells had 616 genes in common with ESCs, signifying a 42% overlap in the genetic profile of the cells. This finding is significant, as these genes play a role in both cellular function, such as cell cycle, RNA post-translation modification, cell death and DNA replication, and stem cell identity, such as self-renewal and pluripotency. Additionally, IPA determined that shared signal-ing pathways for DNA replication, recombination, and repair were significantly enriched between ELA cells, ESCs and iPSCs (right-tailed Fisher’s test, α=0.05), as shown in Table 4. Collec-tively, our data suggests that ELA cells are functional ASCs with a unique set of expressed genes that are not shared with other categories of stem cells.

Immunomodulatory capacity of ELA cells

There was no significant difference between irradiated and non-irradiated ELA cells co-cultured with fresh PBMCs in var-ious ratios. Increasing the number of ELA cells did not influ-ence the proliferative capacity of T cells in the co-culture (Fig-ure 5A). In additional experiments, fresh and cryopreserved ELA cells were co-cultured with freshly isolated PBMCs from three healthy donors at a 1:10 ratio for 5 days (Figure 5B). No significant proliferation of allo-reactive T cells was observed. The SI was 1.72 (± 0.19; n=3) for fresh ELA cells and 1.29 (± 0.36; n=3) for cryopreserved cells. Treatment of ELA cells with the mitogen phytohemagglutinin showed no significant proliferation compared with the very active proliferation with freshly isolated PBMCs (Figure 5B). ELA cells at a ratio of 1:10 elicited a moderate suppressive effect on CD3/CD28-stimu-lated, CFSE-labeled T cells (15.6 ± 2.7%; n=3) compared with a relatively low suppression effect by MSCs (2.8% ± 0.25; n=3)

Jacobs Publishers 9

(Figure 5C). ELA cells at passages 3, 5 and 6 suppressed CD3/CD28-stimulated T lymphocytes, indicating that suppression properties are maintained when ELA cells are expanded in cul-ture, although cells at passage 3 were most effective (Figure 5D).

Next, we wanted to determine if ELA cells promoted the expan-sion of CD4+/CD25+ regulatory T cells. ELA cells were co-cul-tured with freshly isolated PBMCs for a period of 5 d, stained with appropriate antibodies, and analyzed by bi-dimensional FACS analysis. The percentage of CD4+ T cells expressing CD25 expanded 3-fold (10 ± 0.36%; n=3) when co-cultured with ELA cells compared with PBMCs (1.6 ± 0.36%; n=3) (p=0.01) (Figure 5E-F). The percentage of CD8+ T cells expressing CD25

was 3.6 ± 0.36% when cultured with ELA cells (n=3) compared with 1.6 ± 0.36% with PBMCs (n=3). Furthermore, a small pop-ulation (< 1%) of T cells co-cultured with ELA cells expressed PD1 (Figure 5E-F). PD1 is generally associated with exhaus-tion of T cells. We also found a modest increase in CD69, a

surrogate marker of T cell responsiveness to mitogen and an-tigen stimuli, in CD3+ T cells cultured with ELA cells (Figure 5E-F). Taken together, these studies indicate that ELA cells might perform their immunosuppressive functions by inhibit-ing T cell proliferation and expanding CD4+/CD25+ regulatory T cells.

MSCs are known to inhibit the expression of activating recep-tors on the surface of NK cells and potentially impair their cy-

Jacobs Publishers 10

Table 4. Common genes expressed in ELA, uESC, IPS, and MSCs together with enriched canonical pathways

Genes expressed in ELA cells were compared with publically available datasets from the NIH Gene Expression Omnibus: The Ingenuity Pathway Analysis (IPA) was used for idenEfying canonical pathways that were significantly enriched (Right-‐tailed Fisher test with a = 0.05)

ELA$vs.$uESC$Common$Genes$ ELA$vs.$IPS$Common$Genes$ ELA$vs.$MSC$Common$Genes$

Top$Molecular$and$Cellular$Func;ons$ Top$Molecular$and$Cellular$Func;ons!$

Top$Molecular$and$Cellular$Func;ons!

! !Cell!cycle!! !RNA!Post.Transcrip5onal!Modifica5on!! !Cell!Death!! !DNA!replica5on,!Recombina5on!and!Repair!! !Cellular!Assembly!and!Organiza5on!

! !DNA!replica5on,!Recombina5on!and!Repair!! !Cell!cycle!! !Cellular!Assembly!and!Organiza5on!! !Cell!Death!! !Cellular!growth!and!prolifera5on!

!

! !Cellular!Movement!! !Cellular!Growth!and!Prolifera5on!! !Cell!Death!! !Cell!Morphology!! !Cellular!Development!

Top$Physiological$System$Development$and$Func;on$

Top$Physiological$System$Development$and$Func;on!$

Top$Physiological$System$Development$and$Func;on!

! !Connec5ve!Tissue!Development!and!Func5on!! !Cell.Mediated!Immune!Response!! !Humoral!Immune!Response!! !Cardiovascular!System!Development!and!Func5on!! !Hepa5c!System!Development!and!Func5on!

! !Connec5ve!Tissue!Development!and!Func5on!! !Cell.Mediated!Immune!Response!! !Humoral!Immune!Response!! !Organismal!survival!! !Tumor!Morphology!!

! !Organismal!Development!! !Tissue!Development!! !Cardiovascular!System!Development!and!Func5on!! !!Connec5ve!Tissue!Development!and!Func5on!! !!Skeletal!and!Muscular!System!Development!and!Func5on!

Top$Canonical$Pathways$$ Top$Canonical$Pathways$! Top$Canonical$Pathways$!

! !Cell!Cycle:!!G2/M!DNA!Damage!Checkpoint!Regula5on!! !Pyrimidine!Metabolism!! !Wnt/B.Catenine!Signaling!! !Tight!Junc5on!Signaling!! !Role!of!BRCA1!in!DNA!Damage!Response!

! !Parkinson!Signaling!! !Tight!Junc5on!Signaling!! !Pyrimidine!Metabolism!! !Role!of!BRCA1!in!DNA!Damage!Response!! !Cell!Cycle:!!G2/M!DNA!Damage!Checkpoint!Regula5on!!

! !Hepa5c!Fibrosis/!Hepa5c!Stellate!Cell!Ac5va5on!! !An5gen!Presenta5on!Pathway!! !Oncosta5n!M!Signaling!! !IL.6!Signaling!! !TREM1!Signaling!

Top$Networks$ Top$Networks$ Top$Networks!

! !Cell!Death,!Cancer,!Cellular!Movement!! !DNA!Replica5on,!Recombina5on,!and!Repair,!Cell!Cycle,!Cancer!! !Gene!Expression,!Cell!Cycle,!and!Cancer!! !Cell!Cycle,!Cancer,!Cellular!Growth!and!Prolifera5on!! !!

! DNA!Replica5on,!Recombina5on,!and!Repair,!Cell!Cycle,!Cancer!! !DNA!Replica5on,!Recombina5on,!and!Repair,!Cell!Cycle,!Cancer!! DNA!Replica5on,!Recombina5on,!and!Repair,!Cell!Cycle,!Cellular!Assembly!and!Organiza5on!! !Cellular!Assembly!and!Organiza5on,!RNA!Post.!Transcrip5onal!Modifica5on,!Cardiovascular!Disease!! Gene!Expression,!Cell!Cycle,!Dermatological!disease!and!condi5on!!!!

! !Infec5on!Mechanism,!Cancer,!Gene!Expression!! Cellular!Movement,!Cancer,!Cell.to.Cell!Signaling!and!Interac5on.!! !Cellular!Growth!and!Prolifera5on,!Cancer,!Cell!Cycle!! !Gene!Expression,!Cell!Death!,!Tissue!development!! !Gene!Expression,!Developmental!Disorder,!Gene5c!Disorder!

totoxic activity. To evaluate potential ELA cell-mediated inhi-bition of NK cell lytic potential, we performed cytolytic assays. Purified populations of NK cells were co-cultured overnight with and without ELA cells at a 1:1 ratio and than exposed to [51] Cr-labeled K562 target cells at various ratios. Cytolytic ac-tivity was measured by [51] Cr release. NK cells pre-incubated with ELA cells demonstrated >60% reduction in cytotoxic ef-fects. This reduction was consistent across decreasing concen-trations of NK cells (Figure 5G). These data suggested that ELA cells have a suppressive effect on NK cells.

Figure 5. Immunomodulatory potential of early lineage adult (ELA) cells. (A) Stimulation of allo-reactive T cells was determined by co-culturing irradiated and non-irradiated ELA cells with fresh-ly isolated peripheral blood mononuclear cells (PBMCs) at ratios of 1:10, 1:100 and 1:1000 in triplicate for 5 d. PBMCs were isolated from healthy donors, and previously expanded ELA cells were irradiated. To determine the proliferation of allo-reactive T cells, cultures were pulsed with 3[H]-Thymidine (1 μCi/well) 18 h prior to harvesting. Control background Thymidine uptake in PBMCs alone was (487 ± 62.9; n=5). Bar graphs represent the mean ± SEM of 5 separate exper-iments. (B) The effectiveness of cryopreserved ELA cells to stimulate allo-reactive T cell proliferation was determined as in (A). Bar graphs represent the mean of 3 separate experiments ± SEM. (C) Immuno-suppressive properties were further investigated by culturing ELA cells or MSCs with 5-6-carboxyfluorescein diacetate succinidyl ester (CFSE)-labeled allo-reactive T cells at various ratios in triplicate. Af-

ter 5 d culture, T cells were analyzed by flow cytometry to determine CFSE fluorescence. T cell suppression was expressed as the Prolifer-ation Index. (D) The immunosuppressive effectiveness of each ELA cell passage was determined by using different passage numbers of cells co-cultured with CFSE-labeled T cells. (E) A bivariate dot plot analysis of a representative experiment of expanded ELA cell/freshly isolated PBMC co-culture (1:10 ratio) to determine the expression of CD25 on CD4+ and CD8+ T cells and the expression of CD69 or PD1 on CD3+ T cells (* p < 0.001; ** p < 0.01 as compared with MSCs). (F) Bar graphs represent the mean of 3 separate experiments ± SEM (* p < 0.001 as compared with PBMCs alone). (G) The inhibitory effect of ELA cells on NK cell cytotoxicity was demonstrated by co-culturing NK cells with or without ELA cells prior to incubation with target cells at various ratios.

Discussion

Many groups have studied human BM stromal cells and demon-strated phenotypic and functional heterogeneity [22,31,32]. It has been hypothesized that naïve stem cells have a greater ca-pacity to differentiate into functional cells and tissue [33]. Stem cell nomenclature used to refer to a heterogeneous population of marrow stromal cells, but it now just refers to MSCs, a sin-gle type of ASCs. During early bone marrow studies, cells with self-renewing capacity were discovered and referred to as stem cells. These stem cells were later subdivided into hematopoi-etic and non-hematopoietic populations [34,35]. HSCs, one of earliest and best characterized BM stromal stem cell types, al-lowed the first truly successful cellular therapies. To date, over 50,000 HSC transplants have been performed worldwide [36]. Non-hematopoietic MSCs were commonly identified by their ability to differentiate into mesodermal tissues [37]. However, not all marrow stromal cells are phenotypically or function-ally identical, and therefore not all are capable of differenti-ating into similar tissues [38-40]. Most of our understanding of marrow stromal cells comes from studies of MSCs, leading many to believe that there is one category of ASC derived from the bone marrow stroma that has a hierarchical relationship with other ASCs [41]. However, within the bone marrow stro-ma there exists a heterogeneous population of ASCs, such as mesenchymal precursor cells (MPCs), marrow-isolated adult multilineage inducible (MIAMI) cells [42], multipotent adult progenitor cells (MAPCs) [43], and very small embryonic-like (VSEL) stem cells [44]. Recently, our group isolated ELA cells, a population of ASCs that contribute to the heterogeneity of the marrow stromal cell population. Despite standardization of protocols for stem cell biomanufacturing, many protocols start with a heterogeneous population that might not repre-sent the appropriate stem cell for a particular treatment. To address this concern, we plan to focus our future studies on identifying subsets of the primitive stem cells based on mo-lecular profiles with the intention of clonally expanding them.

ELA cells represent a CD99+/MUC1+/CD235a-/MHC class I-


population from the SF of osteoarthritic patients. The prop-erties of these cells were determined by protein and mRNA analysis for pluripotency genes and proteins, such as OCT4 (embryonic form), REX1, and NANOG. Our isolation method was less cumbersome than those required for the enrichment of stem cells from other tissues. The primary reason for this was the absence of RBCs in the SF. Accurate measurements of ELA cell size, volume, and viability were performed with the CASY Cell Counter and Analyzer system. ELA diameter was 4-6 μm, in contrast to RBCs, which have a diameter of 6-8 μm. No-tably, the majority of previous stem cell studies have utilized forward and side scatter perimeters on flow cytometry, which excludes cells < 6 μm. This gating strategy may explain why the ELA cell population has been overlooked until now.

Our studies have shown that the genes ELA cells share with MSCs are not the same as those shared with IPS cells or ESCs. ELA cells are similar in size to VSEL cells, which express OCT-4 and can differentiate into the three germ layers [44]. However, the ELA cell population does not express CXCR4, SSEA-4, CD34, or CD133, which are additional markers used to identify the VSEL cell population [44]. Moreover, ELA cells proliferate in the absence of feeder cells, making them functionally distinct from VSEL cells. Taken together, these findings suggest that the ELA cell population is distinct from the VSEL cell popula-tion. This raises the question of whether ELA cells represent a heterogeneous population of primitive stem cells and whether these subsets can be isolated and clonally expanded for thera-peutic strategies for tissue repair.

In addition to the expression of the pluripotency gene tran-scripts NANOG, OCT4, REX-2, and DPPA3, ELA cells also ex-press high levels of MUC1. Our transcriptome studies con-firmed high expression of mucin genes. Recent studies have shown a relationship between MUC1 and ESC differentiation [45]. There is a need to distinguish the most primitive ASCs, in particular ELA cells, from more mature forms, as primitive ASCs might possess broader differentiation capacity [33] and function as a better starting population for stem cell therapy. Future studies will involve the use of cell surface proteins such as MUC1 to distinguish between primitive and mature ASCs. Collectively, our findings suggest that ELA cells might repre-sent a primitive form of ASCs, making them useful for stem-cell-based treatment.

Although ELA cells reside in a dormant state in the SF (unpub-lished data), their origin is unclear. It is unlikely that SF con-verts MSCs into ELA cells. We hypothesize that the origin of ELA cells is the BM and not the systemic circulation. We base this hypothesis on elegant studies performed by Nakagawa and colleagues, who demonstrated that BM stromal cells migrate directly from the BM to the joint space in a collagen-induced arthritis model [46,47]. We propose that the enhanced migra-tion of ELA cells into the joint space (synovial cavity) increas-

es as a result of inflammation that accompanies osteoarthri-tis. This inflammatory environment is known to increase the number and size of bone canals, which allow communication between the BM and the synovial cavity [46,48]. This increase in bone canal size may allow more ELA cells to migrate into the joint space. Under non-inflammatory conditions, these canals are smaller in size and number, thus limiting the number of ELA cells in the joint space.

Our histochemical data suggests that ELA cells can differenti-ate into various tissues. However, molecular profiling of these tissues showed that transcripts specific for other tissues are also expressed, some of them at high levels. This property of ASCs differentiated in vitro might represent an intermediate phenotype that possesses the ability to transdifferentiate into other tissue types. It is also possible that in vitro culture con-ditions are not representative of in vivo conditions that induce terminal differentiation. These findings support a concept described by Quensenberry and colleagues suggesting that expansion and differentiation of cells and changes in gene ex-pression are continuous and reversible, and that sorting stem cells by static cell surface proteins may leave behind a large percentage of potential stem cells [49].

It was determined that MSCs possess immunomodulatory properties [50,51] and might play specific roles in the main-tenance of peripheral tolerance, transplantation tolerance, au-toimmunity, tumor evasion, and fetal-maternal tolerance [50]. We focused on the role of ELA cells in modulating the immune response by activating T cells. We demonstrated that ELA cells do not induce an allo-immune response, suggesting that they primarily immunomodulate suppression by affecting the ef-fector arm of the immune response. Notably, our in vitro data suggests that ELA cells suppress T cell activation and induce regulatory T cells. Moreover, ELA cells did not upregulate a surrogate marker of T-cell responsiveness (CD69) in CD3+ T cells [52]. ELA cells might interfere with T cell function in a PD-1 independent pathway [45]. Although PD-1 was not up-regulated in either CD4+ or CD8+ T cells, this does not pre-clude the possibility that ELA cells secrete factors or express cell surface proteins that may modulate T cell function. In ad-dition, ELA cells were shown to inhibit the cytolytic capacity of NK cells. Taken together, these findings suggest that ELA cells evade the immune system by interfering with adaptive and in-nate immunity.

Our observations of the ELA cell population are of fundamen-tal importance to the field of regenerative medicine and the de-velopment of cell therapeutics. There is an ever-growing need for stem cells that replace, regenerate, and modulate immune function. However, relying on cell and tissue donation is unre-liable and cannot address the need for ASCs. Biomanufactur-ing of ASC therapies is the most logical option [53]. Although ASCs can be efficiently expanded in the laboratory, this expan-


sion cannot be easily translated to large-scale production for therapeutic purposes due to technical issues. To address these concerns, further studies are underway to assess whether pro-longed culture periods affect the expression of ELA cell-specif-ic genes and ELA cell function.

Conclusion

We have demonstrated the existence of a population of ASCs, referred to as ELA stem cells, which reside within the SF of osteoarthritic joints. These cells are phenotypically distin-guishable from other ASCs. ELA cells are also molecularly dis-tinguishable from ESCs, IPS cells, and MSCs by their unique gene expression patterns. ELA cells represent a new form of primitive ASCs with in vitro tissue regenerative capability that might benefit regenerative therapies. As with MSCs, which are a promising tool for therapeutic approaches aimed at inhibi-tion of the immune response, ELA cells might also be useful for preventing or suppressing graft-versus-host disease in patients undergoing allogeneic HSC transplantation or for the treatment of certain autoimmune diseases. In conclusion, ELA cells have far reaching implications in both basic research and regenerative medicine.

Acknowledgements

The authors gratefully acknowledge the helpful advice provided by Drs. Andrew Makarovskiy and John Garvey. In addition, we gratefully acknowledge the help and assistance provided by the logistics staff for collecting and preparing synovial fluid samples for research purposes. Finally, we would like to acknowledge the patients who participated in providing samples for the studies.

Funding

Research reported in this publication was supported by the Department of Orthopedic Surgery at Brigham and Women’s Hospital and Asclepius Laboratories. The funders had no role in study design, data collection and analysis, decision to pub-lish, or preparation of the manuscript.

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