A Protocol for the Isolation,Culture,and Cryopreservation of Umbilical Cord-Derived Canine Mesenchymal Stromal Cells: Role of Cell Attachment in Long-Term Maintenance

Abstract

Mesenchymal stromal cells (MSCs) hold great promise in the field of regenerative medicine due to their ability to create a variable localized anti-inflammatory effect in injuries such as Crohn's disease and osteoarthritis or by incorporation in tissue engineered constructs. Currently, the MSC literature uses rodents for preclinical disease models. There is growing interest in using naturally occurring disease in large animals for modeling human disease. By review of the canine MSCs literature, it appears that canine MSCs can be difficult to maintain in culture for extended passages and this greatly varies between tissue sources, compared with human and rodent MSCs, and limited lifespan is an obstacle for preclinical investigation and therapeutic use. Research using canine MSCs has been focused on cells derived from bone marrow or adipose tissue, and the differences in manufacturing MSCs between laboratories are problematic due to lack of standardization. To address these issues, here, a stepwise process was used to optimize canine MSCs isolation, expansion, and cryopreservation utilizing canine umbilical cord-derived MSCs. The culture protocol utilizes coating of tissue culture surfaces that increases cellular adherence, increases colony-forming units-fibroblast efficiency, and decreases population doubling times. Canine MSCs isolated with our protocol could be maintained longer than published canine MSCs methods before senescing. Our improved cryopreservation protocols produce on average >90% viable MSCs at thaw. These methods enable master-bank and working-bank scenarios for allogeneic MSC testing in naturally occurring disease in dogs.

Introduction

Multipotent mesenchymal stromal cells (MSCs) are a heterogeneous population of cells that includes stem, progenitor, and differentiated cells. MSCs were originally described in the 1960s as the fibroblastic population of adherent cells derived from bone marrow (BM) that are distinct from the hematopoietic cells found in that niche [1].

Since then, MSCs have been isolated from adult tissues, such as adipose tissue (AT) and BM, and extraembryonic tissues such as the placenta and umbilical cord (UC). Although there may be differences in the MSC populations isolated from each source that could impact their value for regenerative medicine applications, MSCs derived from UC have several advantages over adult tissue sources: UCs are collected nonsurgically from a discarded tissue, they can be collected with no risk or pain to the donor, and they are collected from individuals of a consistent young age [2,3]. Thus, UC-MSCs are a limitless and noncontroversial source of MSCs.

We [4,5] and others [6 –9] have isolated human UC-MSCs, cryogenically banked, and expanded the cells for potential therapeutic applications. For regenerative medicine applications, UC-MSCs might be preferred to adult tissue-derived MSCs due to age-related or disease-related changes that affect MSCs [10 –12]. UC-MSCs are in human clinical testing both in the United States and abroad (clinicaltrials.gov website, accessed March 2019). UC-MSCs have been successfully isolated from veterinary species including swine [13], cattle [14], goats [15], horses [16], and canines [17 –22]. The literature reveals that the efficiency of UC-MSC isolation and expansion differs between species and laboratories. The lack of reproducibility is a source of concern in the MSC field [23,24].

Many factors affect the efficiency to generate MSCs. These factors include isolation method, donor pool, efficiency (cell yield per donor), expansion potential of isolated cells, medium formulation, cell plating density, passaging protocol, cryopreservation efficiency, characterization procedures, senescence, and thawing procedure. Based upon the lack of consistency in manufacturing between laboratories, we suggest that MSC protocols need to be both optimized and standardized.

Many common human genetic disorders and many naturally occurring diseases in humans have canine homologs, adding to the importance of canines as a model for human disease [25]. In addition, canines are an important model species for human anatomical injuries, especially those involving large joints [26]. Canine MSCs have been used as an experimental treatment for orthopedic injuries and show positive clinical results in pilot studies [27 –30]. A recent uncontrolled nonblinded clinical trial in 22 dogs indicated the safety and efficacy of neonatal MSCs in canines with arthritis for a 2-year follow-up period [31]. Canine MSCs are under-represented in the scientific literature compared with human and rodent MSCs [PubMed search February 11, 2019; search terms MSCs and (canine or dog) = 317; MSCs and human = 13,070; MSCs and (rat or mouse) = 8,821].

Here, barriers that impede canine MSC testing are addressed. First, some tissue-specific canine MSCs are slower to expand in culture or undergo senescence around passage 6 [13,17,22,25,28], with some groups surpassing six passages [32 –34]. Second, a consensus set of canine MSC antibodies for characterization has not been established [35]. Having a standardized panel of monoclonal antibodies for cell surface markers CD105, CD44, CD73, CD90, DLA class I and II, CD31, CD45, and CD34 would constitute a minimal set of MSC characterization antibodies. Third, trilineage differentiation has been demonstrated by some groups [17,25,28,36 –41], but not by other groups [19,22,32,42 –46], which suggests that standardized osteogenic, chondrogenic, and adipogenic differentiation protocols for MSCs are needed.

As we provided for human UC-MSCs [4,5], in this study we provide protocols for isolation, expansion, and characterization for canine UC-MSCs. Standardizing protocols may improve the ability to compare results across laboratories and enable clinical translation. Our methods provide healthy viable canine MSCs that can be cryopreserved, thawed, and expanded. The characterization standards for canine MSCs provided here are not comprehensive. However, the protocols provided here remove key barriers and, thus, enable canine UC-MSCs research.

Materials and Methods

UC collection

Institutional animal care and use committee (IACUC) reviewed and approved the current research. The current protocol was deemed an “exempt animal use activity” under guideline no. 21 exemption policy 2.3 “…studies that do not use live animals provided that the tissue is obtained from an IACUC-approved source and is discarded in accordance with all relevant state laws and institutional policies governing disposal of hazardous waste.”

Canine UCs donated after cesarean-section births with owner–informed consent were used. In brief, the dam was anesthetized and placed in lateral recumbence; the abdomen was shaved and surgically prepared for a lateral celiotomy approach to the uterus. The uterus was exteriorized, a single hysterotomy was made, and the pups were delivered. The UCs were separated from the placenta and placed in a sterile transport medium made from an isotonic neutral buffered solution supplemented with 1% antibiotic–antimycotic (Catalog No. 15240062; Gibco) and placed into a 4°C refrigerator. Cords were kept in a styrofoam cooler during transport. Once received in the laboratory, cords were refrigerated at 4°C until processing. All cords were processed within 5 days of birth. Donor breed information was recorded when provided by the clinic, but is not evaluated here.

Gelatin coating of tissue culture plastic

A solution of 0.1% w/w porcine skin gelatin (Catalog No. G2500-100G; Sigma Aldrich) dissolved in distilled water was sterilized and cooled before use. Gelatin solution was added to the flask or well and swirled for 10–15 s. Gelatin was removed and the plate or flask was air dried in the biological safety cabinet (BSC). The dried plates were sealed tightly and stored refrigerated until use, or they were used immediately after drying.

Isolation

UCs were collected from an entire litter and litter size was not considered as a variable. In contrast to human cords, puppies' cords have a complex vascular structure that arborizes making it difficult to determine the vascular material of an individual (boxes in Supplementary Fig. S1). Furthermore, the UCs from littermates were intertwined within the fetal adnexa tissue. The cords were stripped away from the adnexa without stretching or ripping (Supplementary Fig. S1), and pooled, then the length was measured for a desired amount per tube.

Processing of UCs and trimming were performed in the BSC. The cords were rinsed repeatedly in 37°C Dulbecco's phosphate-buffered saline (DPBS) containing 1% antibiotic–antimycotic (1% A/A; Catalog Nos. 14-190-250, 15-240-062; Gibco). The cord length was measured and it was cut into 0.25 cm sections, then transferred into a C-tube (Catalog No. 130-096-334; Miltenyi Biotech) containing 10 mL of 37°C enzyme solution. The length of UC placed in each C-tube was recorded and classified into <30 cm or >30 cm length. The enzyme solution contained 1 mg/mL hyaluronidase (Catalog No. 02151272; MP Biomedicals), 300 U/mL collagenase type I (Catalog No. 17-100-017; Life Technologies), and 300 U of deoxyribonuclease I (Catalog No. D4263-5VL; Sigma Aldrich) in Hank's balanced salt solution (Catalog No. MT21021CM; Corning).

The cord tissue weight was estimated by subtracting the weight of the C-tube with enzyme from the weight of the tube containing the enzyme and the UCs. The C-tubes were processed in a GentleMACS Dissociator (Catalog No. 130-093-235; Miltenyi Biotech) using standard program “C,” once. The C-tubes were centrifuged at 200 g for 5 min, then incubated at 37°C on a Pelco R2 rotator with a 1,051 sample platform at 12 rpm for 3 h. Next, the C-tubes were processed using the GentleMACS Dissociator standard program B, once, and the solution was filtered (100 μm cell strainer, Catalog No. 22-363-549; Fisher Scientific). The filter was washed with an additional 5 mL of DPBS with 1% A/A solution. The cells were pelleted by centrifugation (200 g for 5 min, room temperature), and the supernatant was discarded.

Red blood cells (RBCs) were lysed by resuspending the cells in 0.5 mL of culture medium and addition of 0.5 mL of lysing buffer (Catalog No. R7757-100ML; Hybri-Max, Sigma Aldrich). Cells were mixed by gentle pipetting for 60 s then diluted with 8 mL of DPBS with 1% A/A solution. Cells were pelleted by centrifugation (200 g for 5 min, room temperature), and the supernatant was discarded. Cell pellet was resuspended in 1 mL of 37°C ACB culture medium (recipe provided hereunder).

An aliquot was removed for live/dead cell count using acridine orange/propidium iodide staining solution (Catalog No. CS2-0106-5ML; Nexcelom Bioscience), on a Nexcelom Auto 2000 Cellometer (immune cells, low RBC program). Cells were plated at a density of 20,000—30,000 cells/cm² on the gelatin-coated tissue culture T-75 flasks (Catalog No. 7202000; Corning) in ACB culture medium (ACB consists of Dulbecco's modified Eagle's medium (DMEM, high glucose; Catalog No. 11965092; Gibco) supplemented with 10% fetal bovine serum (FBS) (Catalog No. SH3007103; HyClone, GE Healthcare Life Sciences), 1% antibiotic–antimycotic (Catalog No. 15240096; Gibco), 1% Glutamax (Catalog No. 35-050-061; Gibco), with or without 10 ng/mL basic fibroblast growth factor (bFGF; Catalog No. PHG0264; Gibco).

Culture

After the first passage, canine UC-MSCs were plated at 20,000 cells/cm² on tissue culture plates or flasks (with or without prior gelatin coating) using ACB cell culture medium warmed to 37°C. The cells were grown at 37°C, 5% CO₂, condensing humidity in a Heracell 150i, or Nuaire AutoFlow 4950 incubator. Half of the volume of medium was replaced every 3 days until the cells reached 80%–95% confluency before passage.

Canine UC-MSCs were lifted with either TrypLE express (Catalog No. 12605028; Gibco) or 1.75% nattokinase (Catalog No. NATT100; Bulk Supplements) for 30 min at 37°C. Cells were dislodged with gentle tapping to completely remove them from the plate. If few cells remained adherent to the plate, an additional 5 min of incubation was used. Detached MSCs were collected and pelleted by centrifugation (200 g for 5 min at room temperature). The supernatant was discarded and the cell pellet was resuspended in 1 mL of fresh warm ACB medium. A live/dead cell count was performed at passage, and the cells were plated in fresh medium on gelatin-coated plates at a density of 20,000 cells/cm², cryopreserved, or discarded. Population doubling time was calculated using the following formula: $\begin{matrix} P o p u l a t i o n d o u b l i n g t i m e = \\ \frac{d u r a t i o n o f c u l t u r e (d a y s) \times l o g (2)}{l o g (f i n a l c e l l c o u n t) - l o g (i n i t i a l c e l l c o u n t)} \end{matrix}$

Cryopreservation

To cryopreserve, UC-MSCs were suspended in 1:1 v/v ratio of ACB cell culture medium and freezing medium (Human Embryonic Stem Cell Cryopreservative, MTI-GlobalStem) at 0°C. MSCs were kept ice cold and immediately transferred to a controlled rate freezing apparatus (Mr. Frosty) and then onto the bottom shelf of a −80°C freezer. After 24 h, the vials were moved to the vapor phase of liquid nitrogen tank for long-term storage.

Trilineage differentiation

UC-MSCs between passages 6 and 10 were differentiated to chondrogenic, osteogenic, and adipogenic lineages after testing 3 different media for chondrogenic, 3 different media for adipogenic, and 2 different media for osteogenic lineages (Supplementary Figs. S2–S4). The formulations tested are given in Supplementary Table S1.

The formulations used in the article were high-glucose DMEM, 1% antibiotic–antimycotic, 10 ng transforming growth factor beta 1 (TGF-β1) (Catalog No. GF346; Sigma Aldrich), 1% FBS, 100 nM dexamethasone (Catalog No. D4902; Sigma Aldrich), 1 mM sodium pyruvate (Catalog No. 11360070; Gibco), and 40 μg l-Proline (Catalog No. P8865; Sigma Aldrich) for 21 days for chondrogenesis; high-glucose DMEM, 1% antibiotic–antimycotic, 5% rabbit serum (Catalog No. R9133; Sigma Aldrich), 100 nM dexamethasone, 200 μM indomethacin (Catalog No. I7378; Sigma Aldrich), 10 μM insulin (Catalog No. 12585014; Gibco), and 0.5 mM 3-isobutyl-1-methylxanthine (Catalog No. I7018; Sigma Aldrich) exposure for 14 days for adipogenesis; and StemPro osteogenesis differentiation kit (Catalog No. A1007201; ThermoFisher, StemPro) using manufacturer's protocol for 21 days for osteogenesis.

In brief, 12-well gelatin-coated plates (Catalog No. CC7682-7512; CytoOne) were used to plate UC-MSCs in triplicate at a lineage-specific density. For chondrogenesis, US-MSCs are concentrated to 8 × 10⁶ cells/mL, and a 5 μL droplet is plated in each well that causes the formation of a micromass of MSCs [4]. For osteogenesis, a density of 20,000 UC-MSCs were plated per well, and 76,000 cells per well were used for adipogenesis. Cells remained in culture for 24 to 48 h in standard cell culture medium before changing to differentiation medium. Half the volume of medium was replaced every 3 days.

After 14–21 days of differentiation, medium was removed and cells were washed using DPBS with calcium and magnesium (Catalog No. 14040-133; Gibco). The cells were fixed with freshly prepared 4% paraformaldehyde in 10 mM phosphate buffer (pH 7.4) for 30 min at room temperature, and then triple washed with DPBS. MSCs were stained with Oil Red O to visualize lipid droplets in adipocytes (Catalog No. HT904-8F0Z; Sigma Aldrich), or with Alizarin Red S to detect calcium crystal in osteocytes (Catalog No. A5533-25G; Sigma Aldrich), or with Safranin O to visualize sulfated glycosaminoglycans in chondrocytes (Catalog No. O0625-100G; Sigma Aldrich). After staining, brightfield images were captured using an Evos FL Auto microscope (Life Technologies).

Colony-forming unit-fibroblast assay

Canine UC-MSCs were plated at 50, 100, and 500 cells/cm² in triplicate on gelatin-coated 6-well tissue culture plates (Catalog No. CC7682-7506; CytoOne) in ACB medium and incubated undisturbed for at least 3 days. The medium was changed every 3 days. Cells grew 10–14 days before fixation with ice-cold 100% methanol for 15 min. Methanol was removed and the plates were washed twice for 5 min with room temperature, sterile Sorenson's phosphate buffered saline (pH 7.4). The colonies were then stained with 1% w/v aqueous methylene blue for 20 min at room temperature, gently washed three times with distilled water, and air dried overnight.

The colonies were counted manually using 4–10 × magnification. Colonies were defined as “clonal” groups consisting of >10 cells. The number of colonies per well was averaged from the technical triplicates at each plating density. By dividing the number of cells plated by the averaged number of colonies, the colony forming efficiency was calculated.

Flow cytometry

Flow cytometry analysis of canine UC-MSCs was adapted from the human UC-MSC protocol we have previously published [5]. All antibody clones were previously tested for their use in immunophenotyping canine MSCs (Supplementary Table S2) or are in routine use by the Kansas State University Veterinary Diagnostic Flow Cytometry core for canines.

In brief, UC-MSCs were cultured until 90%–95% confluent, passaged using 1.75% nattokinase, and reconstituted in blocking buffer containing 1% bovine serum albumin (BSA). An aliquot was removed for viability and cell count, whereas the remainder of the cells were incubated in blocking buffer for 15 min at 4°C. Canine UC-MSCs were centrifuged (200 g for 5 min at room temperature) and reconstituted in 1% BSA with the addition of primary antibody at a dilution of 1:100 (Table 1). Canine UC-MSCs were incubated for 1 h at 4°C protected from light, washed, and centrifuged. If antibody was unconjugated, then secondary antibody was added to 1% BSA at a concentration of 2 μg/mL and incubated at 4°C for 30 min protected from light. Finally, cells were washed with 1% BSA and resuspended in 500 μL of 1% BSA and stored at 4°C protected from light until ran on cytometer.

Table 1.

Antibodies Tested in Flow Cytometry

Antibody	Clone	Fluorophore	Catalog no.
CD5	YKIX322.3	PerCP-eFluor 710	46-5050-42
CD11b	M1170	V450	560456
CD14	TUK4	Alexa Fluor^® 700	MCA1568A700
CD21	CA2.1D6	Alexa Fluor 647	MCA1781A647
CD34	1H6	PE	12-0340-42
CD44	IM7	BV786	563736
CD45	YKIX716.13	Alexa Fluor 488	MCA1042F
CD73	7G2	Purified	41-0200
CD90	5E10	PE-Cy7	25-0909-42
CD90	YKIX337.217	APC	12-5900-42
CD105	OTI8A1	Purified	AB156756
HLA-DR	L243	BV650	307602
Goat antimouse IgG H + L	Polyclonal	Alexa Fluor 488	A-11001

DNA staining was based on published protocols [44,45]. In brief, canine UC-MSCs were cultured and passaged for flow cytometry analysis as previously described, then fixed in glacial ethanol for 1 h at −20°C. UC-MSCs were washed twice with DPBS. Then, 50 μg/mL of RNAse H (Catalog No. EN0201; Thermo Scientific) in DPBS was added to canine UC-MSCs for 30 min at 37°C. RNAse was removed by aspirating the supernatant after pelleting the cells using low-speed centrifugation (200 g for 5 min at room temperature). The cell pellet was washed with DPBS, and the UC-MSCs were resuspended in DPBS containing 50 μg/mL of propidium iodide (PI) (Catalog No. P3566; Invitrogen). The cells were stored at 4°C protected from light until they were analyzed on the cytometer.

The samples were analyzed on a BD LSR Fortessa X-20 SORP flow cytometer (BD Biosciences, San Jose, CA) equipped with 405, 488, 561, and 633 nm lasers and appropriate filters to detect all fluorophores listed in Table 1. Data were acquired, recorded, and analyzed utilizing BD FACSDiva 8.0 software (BD Biosciences). For multicolor labeled samples, the compensation matrix was calculated by the software from individually labeled UltraComp eBeads (ThermoFischer Scientific, Waltham, MA) and applied to the cell sample before data acquisition. Cells were identified by forward and side scatter properties and used as the primary gate to exclude debris and doublets. Unstained cells established background fluorescence and acted as a negative control for cell surface markers. At least 10,000 gated data points were recorded for all samples. Results were generated by overlaying the appropriate fluorescence channel of unlabeled and labeled cell samples.

For DNA analysis, cells were identified by setting the detection threshold to be based off fluorescence in a 610/20 bandpass filter, which would identify only PI stained cells. Positively identified cells were gated using pulse geometry to not only exclude doublets but also reveal the presence, if any, of cells with aberrant DNA content. A histogram was generated from the aforementioned gate for cell cycle analysis.

Reverse transcriptase-polymerase chain reaction

Reverse transcriptase-polymerase chain reaction (RT-PCR) was conducted as previously described [47]. In brief, total RNA was isolated using an RNeasy kit (Qiagen). RNA was treated with DNase before storage and measured using a NanoDrop spectrophotometer. Complementary DNA was synthesized from total RNA using Superscript III First-Strand Synthesis Supermix kit (Invitrogen) primed with oligo-dT 12–18 per the manufacturer's protocol. PCR was performed using a BioRad iCycler: initial denaturation at 95°C for 3 min, 30 cycles of (94°C for 1 min, 53°C–55°C for 30 s, and 72°C for 30 s), and the final extension at 72°C for 10 min. After PCR, the products were resolved on a 1%–2% agarose gel with 100 bp DNA ladder and imaged using ethidium bromine. Primer sequences and amplicon size are listed in Table 2.

Table 2.

Reverse Transcriptase-Polymerase Chain Reaction Primers

Gene	Primer sequence	Product length (bp)	Tm(˚C)	Accession no.
CD34-1	GTGCCAACCTCCACAGAAAT	409	55.6	NM_001003341
CD34-1	TGATGGTACTTGGGGTGTCA	409	55.8	NM_001003341
CD34-2	CCCTTTGGGTTCACAAACAC	356	54.5	NM_001003341
CD34-2	TCCGAACCATTTCCAGGTAG	356	54.2	NM_001003341
CD45-1	CCATACAACTGCTCCCACAA	438	55.0	XM_005622278
CD45-1	ACAAAGCCTTCCCATTCAAA	438	52.7	XM_005622278
CD45-2	AACAGCACTGTTGCCCTTCT	386	57.3	XM_005622278
CD45-2	TGGTCACAATTCACGGTATCA	386	54.0	XM_005622278
CD73-1	ACTGGGACACTCTGGTTTCG	422	56.9	XM_532221
CD73-1	ATTCCTTAAAGCGGCAGGAT	422	54.4	XM_532221
CD73-2	TGCATTGCAGCCTGAAGTAG	434	55.6	XM_532221
CD73-2	CTGTTTTCCCCAATTCCTGA	434	52.7	XM_532221
CD90-1	ACATGTGAACTCCGGCTCTC	420	57.0	NM_001287129
CD90-1	AGAAGCGACTCTGGGACAAA	420	56.2	NM_001287129
CD90-2	CGTGATCTATGGCACTGTGG	440	55.6	NM_001287129
CD90-2	GCAGCACTGGGATTCCTTAG	440	55.7	NM_001287129
CD105-1	AGGAGTCAACACCACGGAAC	424	57.2	XM_005625330
CD105-1	GATCTGCATGTTGTGGTTGG	424	54.3	XM_005625330
CD105-2	CCAATGCTACCGTGGAAGTT	378	55.3	XM_005625330
CD105-2	GATTGCAGAAGGACGGTGAT	378	55.0	XM_005625330

For sequencing, the amplicon of the anticipated size from a randomly selected MSC line was cut from the agarose gel (eg, CD34 409 bp, CD34 356 bp, CD73 422 bp, CD73 434 bp, CD90 440 bp, CD90 420 bp, CD105 424 bp, and CD105 378 bp), and the DNA was extracted and purified. Next, the DNA was cloned into a plasmid and expanded. After expansion, five to eight clones were selected and plasmid DNA was isolated and submitted to the KSU Integrated Genomics core for Illumina sequencing. The DNA sequences were checked for quality before alignment. The DNA sequences were aligned with canine sequences in PubMed.

Statistics

After validating that the analysis of variance (ANOVA) assumptions were met, it was used to evaluate significant main effects and/or interactions. After finding significant ANOVA terms, post hoc testing of planned comparisons was performed using either the Bonferroni correction or the Holm–Sidak method. Those data are presented as average ±1 standard deviation (SD). For pairwise comparisons, Student's t-test was used after confirmation of statistical assumptions. If the ANOVA assumptions were not met, then Kruskal–Wallis ANOVA on ranks was used. Those data are presented in box and whisker plots showing median and 25th and 75th percentile in the box and whiskers showing 10th and 90th percentile, with potential outliers indicated by circles. In text, the data are presented as average ±1 SD unless stated otherwise.

Regression analysis was conducted using Sigma Plot v12.5, and significant relationships were reported (regression line is plotted in cases of significant relationship). In one case, regression analysis indicated nonsignificant trends, and was indicated in text (no regression line is shown in graph). Throughout this article, the entire data set was used, and it included potential outliers. The original data set is available. SigmaPlot version12.5 (Systat Software, Inc.) was used for statistics and generating graphs. The graphs created using SigmaPlot were saved as EMF files. These EMF files were labeled and edited for clarity using ACD Systems of America's Canvas (version 15.5, build 1770) and rendered in TIFF format. In all cases, hypothesis testing was two-tailed and P < 0.05 was considered “significant.”

Results

Isolation of MSCs from canine UC

MSCs were isolated from UCs from 30 litters of pups. The schematic for processing canine UCs is shown in Fig. 1. When necessary the fetal placenta was trimmed away from the UC before processing (Supplementary Fig. S1). Two isolation methods were compared: the explant method and the dissociation method (Fig. 2A). The explant method involves mincing the UC into 0.2–0.5 mm pieces and adhering those pieces to the plastic plate before adding medium. The explant method produced the lowest cell yields but produced >90% viable cells (open circle shown in Fig. 2A, B). MSCs from the explant method did not expand after attaching to the culture plate. This method was not tested further, and the remaining UC from 29 litters were mechanically and enzymatic disrupted before culturing, as we described previously for human UC [4,5].

FIG. 1.

A schematic of the isolation procedure showing the major steps involved for obtaining MSCs from the canine UC. After cesarian section delivery, the cord is removed from each puppy and placed into storage solution. The noncord tissues are dissected and discarded using sterile technique. The length of the cord material is measured (a) and it is ligated into lengths (b), rinsed (c), and minced briefly. A fixed length is added to a Milliteny C-tube with enzymes (d). The tube is processed using a standard program (e) and the tube is incubated before cell pellet isolation, counting, and plating. MSC, mesenchymal stromal cells; UC, umbilical cord.

FIG. 2.

Isolation efficiency from UC. (A) Total cell yields from three different variables tested: explant methods (n = 1, white circle), mechanical and enzymatic extraction of <30 cm of cord tissue (red circles) or >30 cm of cord tissue (black circles). Note that over the course of this experiment, our isolation efficiency improved statistically (positive and significant regression line). Note that the explant method was too inefficient to consider for scale-up. Note that adding more tissue to the C-tube did not improve yield or cell viability. This suggests that scale-up might be by using more C-tubes. (B) Cell viability after isolation. Note that the explant method produced the highest cell viability. This was offset by the lowest yield (in A). In contrast, the mechanical and enzymatic methods produced a viability of between 50% and 80%.

To evaluate the effect of tissue volume on cell yield and viability, either <30 cm of UC (black circles shown in Fig. 2A, B) or >30 cm of UC was loaded into the disruptor (red circles shown in Fig. 2A, B). The isolation yield and viability were not significantly changed by tissue volume. Although there was a significant trend for cell yield at isolation to increase over the course of this study (regression line shown in Fig. 2A), there was no significant trend in cell viability (Fig. 2B).

Stage 1

The first 10 canine UC-MSC lines were isolated and expanded using previously described protocols (labeled Stage 1 in Fig. 3). Specifically, the MSCs were plated on tissue culture plastic, exposed to 10% FBS containing DMEM and other standard supplements, and passaged using 0.025% trypsin-ethylenediaminetetraacetic acid (EDTA). Using these methods, only 1 MSC line of the 10 could be expanded beyond 20 cumulative population doublings (CPDs), 50% of the MSC lines were able to expand beyond 10 CPDs, and only 1 line of 10 was able to be expanded to 15 passages (Fig. 3B, C).

FIG. 3.

Stepwise improvement of canine MSC expansion. (A) Left panel: Comparison of the three stages for cell yield at isolation. Horizontal bar indicates 3E6 cells, as a criterion for suitable starting number for expansion. Over the course of the project our methods improved as evidenced by increasing cell yield over time, indicated by a significant positive regression line. Note also that the three stages, indicated by color code, roughly group and follow the trend line. Middle panel: Ultimate passage achieved for 30 canine MSC lines by stage. Horizontal bar indicates eight passages, as a criterion for minimum expansion capacity of MSCs. Note that at each stage of the project there is a significant increase in the number of ultimate passages achieved per cell line. Right panel: CPDs achieved by 30 MSC lines. Horizontal bar indicates 20 CPDs, as a criterion for minimum expansion of MSCs. Note that positive and significant trend line indicating improvement of MSC manufacturing over project. (B) CPDs for all cell lines versus days in culture for all 30 MSC lines by sStage (Stage 1, left panel; Stage 2, middle panel; Stage 3, right panel). These graphs give an indication of how rapidly MSCs expanded per passage, and the number of CPDs achieved by Stage. (C) CPDs for all cell lines versus passage for all 30 MSC lines by stage (Stage 1, left panel; Stage 2, middle panel; Stage 3, right panel). Note that 15 passages were set as a maximum number of passages, arbitrarily. Note that over stage, more MSC lines were able to reach the arbitrary maximum of 15 passages. In (B, C) horizontal bar indicates 20 CPDs, as a criterion for minimum expansion of MSCs, and vertical bar indicates 60 days in culture, as a target to try to keep manufacturing times as short as possible. (D) Population doubling time by stage and time in culture (Stage 1, yellow boxes; Stage 2, blue boxes; Stage 3, green boxes). Note that the box plots indicate median and 25th and 75th percentile and the whiskers indicate 10th and 90th percentiles. Each stage is further broken into early and late passages. “Early” is defined as passages 0–4 and “late” is defined as passages 5–10. From this, we can see that moving to Stage 2, the addition of gelatin coating and nattokinase for lifting MSCs, but no change to medium did not greatly change MSC growth rate, but tended to reduce senescence. In contrast, in Stage 3, changing the medium formulation increased MSC growth, especially in the early passage epoch. (E) Population doubling time by stage (Stage 1, yellow boxes; Stage 2, blue boxes; Stage 3, green boxes). When population doubling time was averaged (over all passages by stage), a significant increase in growth rate was observed between Stages 2 and 3. The median growth rate of Stage 3 was ∼2.5 days. (F) Loss of MSC proliferation over passage by stage. Percentage of expanding MSC lines by passage and by stage (Stage 1, yellow dot; Stage 2, blue dots; Stage 3, green triangles). Note that while sustaining MSCs in culture showed a marked improvement over the three stages (in F), just 50% of the lines were able to achieve 15 passages in Stage 3. This suggests that improvements in culture conditions may be identified in future study. Dotted lines in (A) indicate statistically significant, positive regression lines indicate improvement over the project. Box plots in (D, E) indicate median, 25th, and 75th percentiles; whiskers indicate 10th and 90th percentiles (potential outliers indicated by filled circles). *Indicates significant (P < 0.05) using a two-tailed test. CPDs, cumulative population doublings.

In Stage 1, we learned that when compared with human UC-MSCs, canine UC-MSCs require higher plating density to expand (20,000 cells/cm² for canine UC-MSCs vs. 10,000 cells/cm² for human UC-MSCs). In Stage 1, we performed pilot experiments with different agents for lifting MSCs off the tissue culture plate for passage (discussed hereunder).

Stage 2

Two modifications in UC-MSC expansion were made in Stage 2. First, tissue culture plastic was modified by coating the plates with gelatin. Second, nattokinase was used for lifting the MSCs at time of passage [48,49]. In Stage 2, the expansion potential of UC-MSC was improved based upon the following observations: first, a significant positive trend line was found for the total cell yield, the ultimate number of passages reached by MSC lines, and the CPDs achieved (Fig. 3A). Second, 6 of 9 (66%) MSC lines expanded beyond 10 CPDs and 3 of 9 (33%) MSC lines expanded beyond 20 CPDs) (Fig. 3B). Third, 3 of 9 MSCs lines expanded to 15 passages without senescence (Fig. 3B, C).

Stage 3

One modification was made in UC-MSC expansion in Stage 3 compared with Stage 2: a growth supplement, bFGF, was added to the medium (ACB medium) together with plating on gelatin-coated plates [50 –52]. This modification further improved in UC-MSC expansion capability. This enhancement was indicated by better yield at initial isolation (9 out of 11 exceeded 3E6 cells at isolation) (Fig. 3A), and an increase to 9 out of 11 (90%) of the MSC lines expanding beyond 10 CPDs, 5 out of 11 (50%) of the MSC lines expanding beyond 20 CPDs, and 5 of 11 MSC lines expanded to 15 passages (Fig. 3B, C).

The faster growth than Stage 2 was indicated by 5 of 11 MSC lines reaching or surpassing 20 CPDs by 60 days of culture (Fig. 3B), and by significantly faster population doubling time, especially in the first five passages (labeled early in Fig. 3D). The relative efficiency to maintain MSCs by stage of development is shown in Fig. 3F.

Effect of gelatin-coated plates

The effect of gelatin-coated tissue culture plates was apparent during culture based upon cellular morphology (Fig. 4) and cumulatively over passage (Fig. 5), but it did not significantly affect the growth rate of MSCs during the first five passages (comparing the population doubling time between Stages 1 and 2 shown in Fig. 3D), or over entire culture period (comparing Stages 1 and 2 shown in Fig. 3E).

FIG. 4.

Effect of gelatin coating of plates on canine MSCs morphology. (A) Canine MSC grown on tissue culture-treated plastic (Stage 1). Note the heterogeneity of the culture and the presence of large flattened cells with stress fibers and debris. (B) Canine MSCs grown on gelatin-coated plastic. Note the more homogeneous appearance of fusiform cells and the smaller cell size. Calibration bar is 1 mm.

FIG. 5.

Effect of gelatin coating of plates on canine MSCs expansion. (A) When comparing canine MSCs from Stage 1 and Stage 2, gelatin coating of the plate tended to improve the CPDs achieved, but this was not statistically significant. (B) In contrast, when comparing the number of passages with senescence for Stages 1 and 2, gelatin-coated plates significantly improved MSC's ability to achieve more passages. (C) Gelatin coating of the tissue culture plastic also improved the ability to perform colony-forming unit-fibroblast assays, as indicated by the number of colonies obtained per number of cells seeded. (D) Improved colony forming efficiency by gelatin coating. Gelatin coating significantly improves number of colonies compared with uncoated plates. Box plots indicate median, 25th, and 75th percentiles; whiskers indicate 10th and 90th percentiles (potential outliers indicated by filled circles). *Indicates significant (P < 0.05) using a two-tailed test.

Subjectively, UC-MSCs grown on gelatin-coated plates appear to experience less stress than MSCs grown on standard tissue culture plates (compare Fig. 4A and B). For example, UC-MSCs grown on tissue culture plastic had more debris, and more of the large flattened cells with stress fibers (Fig. 4A) than MSCs grown on gelatin-coated plates (Fig. 4B). This subjective observation was supported by other observations. As shown in Fig. 5A, there was a trend for MSCs grown on gelatin-coated plates to have greater CPDs, and as shown in Fig. 5B, there were significantly more passages until senescence when MSCs were grown on gelatin-coated plates. The attachment of MSCs to gelatin-coated plates also enhanced the colony forming fibroblast efficiency (Fig. 5C, D). As MSCs grew on gelatin-coated plates, a pattern in the arrangement of their cell bodies was observed when the same field was observed from day to day (Supplementary Fig. S5).

Comparing method of lifting cells for passage

Based upon our experience with human MSCs (data not shown) [4] and based on the canine MSC literature, we assumed that 0.025% trypsin-EDTA was suitable for canine MSCs. To evaluate this hypothesis, pilot experiments were conducted using 0.025% trypsin-EDTA compared with TrypLE, dispase, TrypLE express, and 1.75% nattokinase. We determined that dispase, TrypLE, and Trypsin-EDTA negatively impact yield, viability, and downstream effects (comparison data not shown). Trypsin-EDTA and dispase were not successful and resulted in failure to proliferate after passage (data not shown). TrypLE was unable to efficiently lift canine MSC from the plate for passage (data not shown).

In contrast, nattokinase improved MSC viability at passage compared with TrypLE express (Fig. 6A, B). The difference between cell viability and MSC yield between TrypLE express (0.05%) versus nattokinase (1.75%) was significant (Fig. 6B, D). Nattokinase was used to lift MSCs in Stages 2 and 3.

FIG. 6.

Nattokinase improved MSC passage compared with TrypLE express. (A) Side-by-side comparison of cell viability when MSCs at the same passage were lifted by TrypLE express or nattokinase (83 independent trials). (B) Summarized data from (A). Nattokinase significantly improved MSC viability at passage (median of 98.1% for nattokinase, median of 95.6% for TrypLE express). (C) Side-by-side comparison of MSC yield at passage when MSCs of the same passage were lifted with TrypLE express or nattokinase. (D) Summarized data from (C). Nattokinase significantly improved cell yield at passage (median of 1.99E6 cells for nattokinase, median of 1.64E6 for TrypLE express, a 21.3% improvement). Box plots indicate median, 25th, and 75th percentiles; whiskers indicate 10th and 90th percentiles (potential outliers indicated by filled circles). *Indicates significant (P < 0.05) using a two-tailed test.

Cryopreservation effects and ability to revive UC-MSCs from cryostorage

It appears that canine UC-MSCs are less robust than human UC-MSCs in terms of their expansion potential, response to chemical stress, and attachment ability. We then queried whether canine UC-MSCs were sensitive to cryopreservation and could be revived and expanded after cryostorage. To answer this question, 33 UC-MSC samples were frozen and stored for an average of 185 days (median 191 days, range 4–448 days, 25th percentile 34.5 and 75th percentile 292.5 days) in vapor phase liquid nitrogen then thawed and tested for viability. As expected, there was a significant drop in MSC viability of 7.9% ± 4.8% after thawing (Fig. 7A, B). Although viability decreased due to freeze/thaw, there was no significant effects of length of storage on change in MSC viability (Fig. 7C, D). Note that the regression line shown in Fig. 7D is not significant. All thawed cell lines expanded for multiple passages after thaw.

FIG. 7.

Cryostorage reduces the viability of canine MSCs. (A) Side-by-side comparison of the viability of 33 MSC samples before (black circles) and after cyrostorage (white circles) shows consistent loss of viability resulting from freeze/thaw cycle. (B) Summarized data from (A). As expected, cryostorage significantly reduces canine MSC viability. (C) Side-by-side comparison of the effect of time on MSC viability at thaw. Each vector indicates the change in viability from fresh to after thaw for an individual canine MSC sample. X-axis indicates the number of days in cryostorage before thaw. By inspection, there did not appear to be a time-dependent loss in viability due to cryostorage. (D) Change in viability (indicated by vector length in C) versus days frozen. Regression analysis reveals no significant relationship between change in viability over time (although a negative trend line is observed). Box plots indicate median, 25th, and 75th percentiles; whiskers indicate 10th and 90th percentiles (potential outliers indicated by filled circles). *Indicates significant (P < 0.05) using a two-tailed test.

Trilineage differentiation

Many articles in the canine MSC literature do not demonstrate trilineage differentiation, and for that reason, some have argued that differentiation to two lineages is sufficient to demonstrate multipotent progenitors in canine MSCs. In pilot testing, eight different differentiation methods gleaned from the literature were compared for trilineage differentiation (Supplementary Figs. S2–S4 and Supplementary Table S1). As shown in Fig. 8, canine UC-MSCs successfully differentiate to bone-forming, cartilage-forming, and fat-forming lineages. The efficiency to differentiate canine UC-MSCs to adipocytes was low, but this probably reflected cell loss during differentiation and further optimization is likely possible.

FIG. 8.

Trilineage differentiation of canine MSCs (line 30 at passage 5). (A–C) Negative control (control MSCs): MSCs in normal culture medium and stained to demonstrate background levels of calcium deposition using Alizarin Red S (A, for osteogenic differentiation), acidic proteoglycan staining using Safranin O (B, for chondrogenic differentiation), and lipid droplets using Oil Red O (C, for adipogenic differentiation). (D–F) MSCs after trilineage differentiation (differentiated MSCs). Calcium matrix deposition after 21 days of osteogenic differentiation (D). MSC micromasses form and stain intensely for acidic proteoglycans after 14 days of chondrogenic differentiation (E). Lipid droplets within MSCs after 14 days of adipogenic differentiation (F). In all cases, staining was performed after fixation. Representative wells shown from technical duplicates. Calibration bar is 400 mm in (A, D), 1,000 mm in (B, E), and 200 mm in (C, F).

Immunophenotyping by flow cytometry

We performed extensive testing of antibodies to find those that work best for canine UC-MSCs (Table 1). While representative data are shown in Fig. 9, we validated the staining patterns provided here in two or three different canine UC-MSC lines at passages 5 to 8. Canine UC-MSCs positively labeled for CD90, CD73, CD44, and CD105, as indicated by the mean fluorescence intensity shift in the monomodal population of gated cells, but the size of the positive shifts was not as large as we have seen previously when working with human UC-MSCs [4,5,53]. We wondered whether the smaller shift was due to using a mouse antihuman CD90 (clone 5E10), instead of a canine-specific antibody. To address this concern, we repeated the flow cytometry using a rat anticanine CD90 (clone YKIX337.217).

FIG. 9.

Immunophenotyping of canine UC-MSCs. Representative flow cytometry histograms with unstained cells (light gray) versus antibody labeled sample (dark gray) for two or three canine MSC cell lines, depending on surface marker. Based on the prominent shift in fluorescence intensity for the entire population, the data are interpreted as follows: MSCs were positive for CD34, CD105, CD44, CD73, and CD90. MSCs were negative for CD11b, CD5, HLA-DR (MHC-II), CD21, CD14, and CD45.

As shown in Supplementary Fig. S6, the positive shift in CD90 was not frankly affected by using a canine-specific rat anti-CD90 compared with the mouse human-specific anti-CD90. We conclude that the surface expression of canine UC-MSCs is less than that of human UC-MSCs. Another observation that confused us was that canine UC-MSCs displayed positive labeling for CD34, which is typically considered a hematopoietic stem cell marker. To confirm whether CD34 was expressed by canine MSCs, we performed follow-up experiments that are described in the next section. As expected, canine MSCs had negative labeling for MHC class II, CD11b, CD14, CD21, CD5, and CD45.

Confirmation of CD34 expression by RT-PCR

Since we observed CD34-positive labeling for canine UC-MSCs, we confirmed this by alternative means. Two PCR primer pairs, spanning an intron, were designed for CD73, CD90, CD105, CD34, and CD45 to perform RT-PCR on RNA samples obtained from 12 different canine MSC lines. As shown in Supplementary Fig. S7, canine MSCs express mRNA for CD73, CD90, CD105, and CD34, as indicated by the finding the appropriately sized amplicon after RT-PCR amplification for both primer pairs tested. As shown in Supplementary Fig. S6, both PCR primer sets confirm expression of CD73, CD90, CD105, and CD34 mRNA. However, the 434 bp PCR primer pair for CD73 showed multiple bands, a strong band between 400 and 500 bp of the expected product, and in lanes 2,3, and 11, a second, larger product, suggesting nonspecific amplification.

Therefore, we confirmed the specificity of the PCR findings for CD34, CD73, CD90, and CD105 by sequencing. The sequences were verified to be 99%–100% matches to the expected canine mRNA (Supplementary Table S3). In contrast, canine MSCs did not express mRNA for CD45 to a detectable amount. This suggests that the flow cytometry results are valid since 11 of 12 lines tested (>91%) of the canine UC-MSC lines expanded using our culture, and passage conditions express CD34 mRNA and protein.

Evaluation of cell cycle by flow cytometry

Canine UC-MSC ploidy and cell cycle partitioning were evaluated using flow cytometry, as shown in Fig. 10. The analysis of cell cycle revealed ∼78% of the cells in G0/G1, ∼16% cells in G2/M phase, and ∼8% of the cells in S phase. Note that no tetraploid or aneuploid cells were detected.

FIG. 10.

Flow cytometric analysis of canine MSC cell cycle and polyploidy. The left panel illustrates the gating strategy to exclude doublets and debris. The histogram in the right panel indicates the stage of cell cycle based upon DNA staining. The majority of canine MSCs (∼78%) were in the G0/G1 phase with fewer cells in the G2/M phase (16%), and 6% in the S phase.

Discussion

In this study, canine UC-MSC isolation and manufacturing protocols were generated using UCs from 30 litters. Five new findings encapsulate this study. First, the manufacturing of canine UC-MSCs was fundamentally different from that of human UC-MSCs. By using human UC-MSC manufacturing protocols on canine cells, as done in Stage 1, most lines cease proliferation or senesce rapidly, since only 50% of the MSC lines could be maintained beyond passage 4.

Second, canine UC-MSC manufacturing was improved by adding cell attachment factors, such as gelatin -coating to tissue culture plates. Study by Devireddy et al. had previously indicated that gelatin coating was important for canine MSCs when they are expanded in serum-free defined medium [54]. This study indicates that addition of this attachment factor extends the longevity of MSC culture and improves colony-forming unit-fibroblast efficiency when MSCs are grown with FBS-supplemented medium. When human MSCs are cultured in serum-free conditions, the addition of attachment factors is critical for their expansion [53].

These changes, represented here by shift from Stages 1 to 2, enabled 50% of the MSC lines expand to passage 8. Furthermore, use of gelatin coating significantly enhanced longevity of MSCs' self-renewal capability indicated by higher CPDs, and improved colony forming efficiency.

Third, an improvement was noted when nattokinase was used for lifting MSCs for passage. Nattokinase is a serine protease and a fibrinolytic enzyme isolated from Bacillus subtillis natto B-12 in traditionally fermented Japanese soybean. Previously, Carrion et al. reported that nattokinase could extract BM-derived MSCs out of a fibrin gel better than trypsin-EDTA or TryplLE [49]. We found nattokinase to be more effective for lifting canine MSCs than standard lifting agents, terms of yield, and viability. Lifting MSCs and passaging through trypsin-EDTA, dispase, or TrypLE reduced passage yield and viability. In short, nattokinase significantly improved cell viability and canine UC-MSC yield at passage.

Fourth, canine UC-MSC viability is reduced by cryopreservation, as expected. However, the impact of cryopreservation on canine UC-MSCs is no worse than that on human UC-MSCs (data not shown), and all 33 canine UC-MSC lines tested here re-entered the cell cycle and expanded after cyrostorage. Importantly, the length of cryostorage had no significant impact on revival viability or expansion capability of canine UC-MSCs.

Fifth, we evaluated antibodies to be used as a standardized panel for evaluation of canine UC-MSCs. While overcoming the early senescence of canine MSC was a critical hurdle, the next challenge is to characterize canine MSCs. This is called a challenge because no consensus set of antibodies has been described for canine MSC surface markers. In this study, we addressed this issue, and the monoclonal antibodies used in the canine MSC field were reviewed, and a list of antibodies that have been tested for canine MSC characterization is provided in Supplementary Table S2 [17 –19,22,31,32,37,45,46,54 –65]. Antibodies used here are highlighted on this list.

As can be seen when reviewing this table, the flow cytometry results using these antibodies were conflicting: Some of the antibodies performed well, meaning that they produced consistent results across different laboratories, and others were less consistent. In Table 1, the antibodies that worked in this study, and henceforth might be considered for a standard flow cytometry marker set for canine MSC immunophenotype characterization, are provided.

Two points raised by our flow cytometry study bear additional discussion. First, the positive shift for CD73 and CD90 staining was less than what we have seen in human UC-MSCs [52 –54]. We tested whether using an anticanine-specific CD90 would improve the flow cytometry positive shift, compared with the antihuman CD90. As shown in Supplementary Fig. S6, the positive shift was not frankly different, suggesting that the expression of CD90 was not as high in canine MSCs.

Second, canine UC-MSCs appear to express CD34. This observation is controversial, since it is in contrast to the International Society for Cell and Gene Therapy (ISCT) consensus marker set for human MSCs [35]. However, it should be noted that three other laboratories have reported CD34 staining of canine MSCs, while most laboratories do not observe CD34 staining (Supplementary Table S2). Note also that the flow cytometry results for CD34 were confirmed by RT-PCR, and 11 out of 12 UC-MSC lines expressed CD34 mRNA (shown in Supplementary Fig. S6), and further verified by DNA sequencing of the amplicons (data not shown). Together, this suggests that our cell source, culture, and passaging conditions result in CD34 expression by UC-MSCs. These five new findings indicate the importance for optimizing cell culture conditions and may have downstream impact on the clinical testing of canine UC-MSCs.

In the human MSC field, commercial human MSC flow cytometry characterization kits provide as a “standard” for comparison of MSC surface markers per the ISCT definition across the field. In contrast, no standardized antibody kit is available for the canine to facilitate between laboratory MSC comparisons. Canine MSCs positive surface marker expression, for example, the shift in mean fluorescence intensity, is less robust than that observed in human MSCs. Specifically, the shift in fluorescence intensity for canine MSCs for markers CD73, CD90, CD105, and CD44 is smaller than that observed in human MSCs. This could be due to differences in the affinity of the antibody for canine versus human molecule, or differences in the expression level between species, or due to cell culture-related “artifact.”

Canine UC-MSCs were negative for hematopoietic markers CD45, CD5, CD21, CD14, and CD11b, and for HLA-DR (MHC class II), like human MSCs. Review of the literature provided in Supplementary Table S2 revealed that some laboratories reported CD34-positive canine MSCs [22,42,59,63], and other laboratories, using the same CD34 clone, find no MSCs staining for CD34.

Our review found that CD34-positive cells did not break cleanly such that CD34 expression was found only in tissue-specific MSCs, or only in early passage MSCs. For example, one article found CD34-positive cells in 18.4% AT-derived MSCs and in 3.6% BM-derived MSCs [22]. Another article reported 29.2% CD34-positive AT-derived MSCs [59]. A third article shows that CD34 expression (10%) AT-derived MSCs compared with 1% of the MSCs were CD105 positive. Finally, Ryu et al. indicated that CD34 was expressed at low level in passage 1 AT-derived MSCs, but the expression was lost by passage 7 [63].

Based upon our results and the previous reports, some possible explanations for the disparity in CD34 expression can be considered. First, the observed CD34 staining here is real and CD34 expression in MSCs may be induced by culture or passaging conditions used here, since CD34 antibody clone we used performs consistently with high specificity in canine peripheral blood and BM immunophenotyping (K.K. and N.S., pers. comm.). Flow cytometry results for CD34 are supported by our RT-PCR (shown in Supplementary Fig. S6) and DNA sequencing results (data not shown), which demonstrated CD34 expression in 11 of 12 canine UC-MSC lines.

Second, as indicated by 3D cell culture work, it is possible that cell–substrate and cell–cell interactions may alter surface marker phenotype [66 –68]. Follow-up study is needed to determine whether CD34 expression is due to using UC-MSCs, and to determine whether MSCs from other tissue sources can be induced to express CD34 by altered culture conditions.

Previous reports indicated that expanding canine MSCs was problematic, for example, that canine MSCs cannot proliferate beyond passage 7, and that canine MSCs must be characterized around passage 3 [60]. Some previous studies did not demonstrate trilineage differentiation or they performed RT-PCR to demonstrate gene expression in lieu of flow cytometry. We attribute these “problems,” for example, lack of multi-lineage differentiation, loss of differentiation potential, and to senescence associated with inappropriate cell attachment and passaging conditions. This was supported in our pilot studies (data not shown) where UC-MSCs from Stage 1 failed to differentiate or did so at very low efficiency whereas UC-MSCs from Stages 2 and 3 were able to differentiate at high efficiency.

Canine UC-MSCs have different medium requirements than human UC-MSCs, and that the addition of bFGF to the medium formulation significantly improved population doubling time and longevity in culture such that lines could be maintained beyond passage 11. The addition of bFGF is known to have a mitogenic effect on MSCs of other species [50 –52]. In summary, we demonstrated that bFGF not only improved growth rate, but it also may play a role in preventing UC-MSC senescence and improving differentiation efficiency.

Canine UC-MSCs were capable of differentiation to bone-, cartilage-, and fat-producing cells. Note that trilineage differentiation of canine MSCs is a point wherein laboratories have not consistently demonstrated the ISCT MSC definition. While saying this, we also observed low efficiency to differentiate UC-MSCs to adipocytes.

We suggest that trilineage differentiation be adopted as part of standard for defining canine MSCs to meet the ISCT “multipotent cell” definition. We also suggest that further refinement in differentiation protocols would improve compliance. Note that more recently the ISCT released revised MSC characterization criterion that includes functional bioassays. We consider the addition of a functional bioassay important for canine UC-MSC characterization but it is beyond the scope of the present report.

Canine MSCs derived from different tissues such as AT, BM, or tissues discarded at birth such as placenta, amniotic membranes, or UC have been compared by others. In that regard, the advantages of MSCs derived from UC are clear. For example, MSCs from the UC are isolated from subjects of a consistent young age, and there is the potential to create banks of allogeneic cells for clinical use. Therefore, manufactured and banked UC-MSCs have the potential as an off-the-shelf allogeneic product, similar to UC blood hematopoietic stem cells. In that regard, this report provides an important contribution to the canine MSC literature.

MSCs are widely investigated in the field of regenerative medicine, as indicated by the number of MSC clinical trials listed on the clinicaltrials.gov website. This clinical research “push” is not limited to human medicine, since there have been MSC trials conducted in veterinary medicine, too. Thus, the demand for MSCs in large quantities and of “clinical grade” is present in both human and veterinary medicine. This demand for MSCs, however, is not well met. There is no standardized method to isolate, expand, characterize, and freeze/thaw MSCs that is universally accepted. There is no standard to compare MSC quality for their clinical potency. The lack of such standards might be an impediment for clinical translation, since each laboratory argues its case separately with the FDA. It seems likely that new manufacturing and qualification processes are needed to produce both large numbers but also MSCs manufactured of a quality standard [69].

In conclusion, this report addresses some limitations associated with manufacturing canine MSCs for clinical applications. Although significant improvements are reported here, further optimization might be possible. The knowledge gap between human UC-MSCs and canine UC-MSCs should be closed to maximize the usefulness of this companion animal species as a model for human UC-MSC clinical translation.

Footnotes

Acknowledgments

We thank Drs. Charan Ganta (Kansas State University) and Dori Borjesson (University of California, Davis) for critically reading an earlier draft of this article and providing input. We thank Dr. Kent Law and his staff from Symbioun, Inc. (Abilene, Kansas) as well as Dr. Jeannie Binns and her staff at NEK Veterinary Services Group (Hiawatha, Kansas) for their crucial role in the collection and donation of discarded canine UC tissue. The authors acknowledge the artistic support of Ellen Weiss for producing the graphics in .

Author Disclosure Statement

The authors have nothing to declare. The intellectual property reported here has been disclosed to Kansas State University's Institute for Commercialization and a patent disclosure has been filed.

Funding Information

This study was supported by the State of Kansas to the Midwest Institute for Comparative Stem Cell Biotechnology.

Supplementary Material

Supplementary Figure S1

Supplementary Figure S2

Supplementary Figure S3

Supplementary Figure S4

Supplementary Figure S5

Supplementary Figure S6

Supplementary Figure S7

Supplementary Table S1

Supplementary Table S2

Supplementary Table S3

References

Friedenstein

, Petrakova

, Kurolesova

and Frolova

. (1968). Heterotopic of bone marrow. Analysis of precursor cells for osteogenic and hematopoietic tissues. Transplantation, 6:230–247.

Weiss

and Troyer

. (2006). Stem cells in the umbilical cord. Stem Cell Rev, 2:155–162.

Troyer

and Weiss

. (2008). Wharton's jelly-derived cells are a primitive stromal cell population. Stem Cells, 26:591–599.

Smith

, Cromer

and Weiss

. (2017). Human umbilical cord mesenchymal stromal cell: isolation, expansion, cryopreservation, and characterization. Curr Protoc Stem Cell Biol, 41:1F.18.1–1F.18.23.

Smith

, Pfeifer

, Petry

, Powell

, Delzeit

and Weiss

. (2016). Standardizing umbilical cord mesenchymal stromal cells for translation to clinical use: selection of GMP-compliant medium and a simplified isolation method. Stem Cells Int, 2016:6810980.

Fazzina

, Mariotti

, Procoli

, Fioravanti

, Iudicone

, Scambia

, Pierelli

and Bonanno

. (2015). A new standardized clinical-grade protocol for banking human umbilical cord tissue cells. Transfusion, 55:2864–2873.

Badowski

, Muise

and Harris

. (2014). Mixed effects of long-term frozen storage on cord tissue stem cells. Cytotherapy, 16:1313–1321.

Chatzistamatiou

, Papassavas

, Michalopoulos

, Gamaloutsos

, Mallis

, Gontika

, Panagouli

, Koussoulakos

and Stavropoulos-Giokas

. (2014). Optimizing isolation culture and freezing methods to preserve Wharton's jelly's mesenchymal stem cell (MSC) properties: an MSC banking protocol validation for the Hellenic Cord Blood Bank. Transfusion, 54:3108–3120.

Han ZC. (2009). Umbilical cord mesenchymal stem cells (UC-MSCs): biology, banking and clinical applications. Bull Acad Natl Med, 193:545–547; discussion 547.

10.

Ganguly

, El-Jawhari

, Giannoudis

, Burska

, Ponchel

and Jones

. (2017). Age-related changes in bone mesenchymal stromal cells: a potential impact on osteoporosis and osteoarthritis development. Cell Transplant, 26:1520–1529.

11.

Lepperdinger

. (2011). Inflammation and mesenchymal stem cell aging. Curr Opin Immunol, 23:518–524.

12.

Alt

, Senst

, Murthy

, Slakey

, Dupin

, Chaffin

, Kadowitz

and Izadpanah

. (2012). Aging alters tissue resident mesenchymal stem cell properties. Stem Cell Res, 8:215–225.

13.

Carlin

, Davis

, Weiss

, Schultz

and Troyer

. (2006). Expression of early transcription factors Oct-4, Sox-2 and Nanog by porcine umbilical cord (PUC) matrix cells. Reprod Biol Endocrinol, 4:8.

14.

Raoufi

, Tajik

, Dehghan

, Eini

and Barin

. (2011). Isolation and differentiation of mesenchymal stem cells from bovine umbilical cord blood. Reprod Domest Anim, 46:95–99.

15.

Kumar

, Agarwal

, Das

, Mili

, Madhusoodan

, Kumar

and Bag

. (2016). Isolation and characterization of mesenchymal stem cells from caprine umbilical cord tissue matrix. Tissue Cell, 48:653–658.

16.

Cremonesi

, Violini

, Lange Consiglio

, Ramelli

, Ranzenigo

and Mariani

. (2008). Isolation, in vitro culture and characterization of foal umbilical cord stem cells at birth. Vet Res Commun 32 Suppl, 1:S139–S142.

17.

Saulnier

, Loriau

, Febre

, Robert

, Rakic

, Bonte

, Buff

and Maddens

. (2016). Canine placenta: a promising potential source of highly proliferative and immunomodulatory mesenchymal stromal cells?. Vet Immunol Immunopathol, 171:47–55.

18.

Seo

, Park

and Kang

. (2012). Isolation and characterization of canine Wharton's jelly-derived mesenchymal stem cells. Cell Transplant, 21:1493–1502.

19.

Ryu

, Kang

, Park

, Kim

, Sung

, Woo

, Kim

and Kweon

. (2012). Comparison of mesenchymal stem cells derived from fat, bone marrow, Wharton's jelly, and umbilical cord blood for treating spinal cord injuries in dogs. J Vet Med Sci, 74:1617–1630.

20.

Groza

, Pop

, Cenariu

, Ciupe

and Pall

. (2016). Canine Wharton's jelly derived mesenchymal stem cell isolation. Agric Agric Sci Procedia, 10:408–411.

21.

Clark

, Kol

, Shahbenderian

, Granick

, Walker

and Borjesson

. (2016). Canine and equine mesenchymal stem cells grown in serum free media have altered immunophenotype. Stem Cell Rev, 12:245–256.

22.

Russell

, Chow

, Dukoff

, Gibson

, LaMarre

, Betts

and Koch

. (2016). Characterization and immunomodulatory effects of canine adipose tissue- and bone marrow-derived mesenchymal stromal cells. PLoS One, 11:e0167442.

23.

Viswanathan

, Keating

, Deans

, Hematti

, Prockop

, Stroncek

, Stacey

, Weiss

, Mason

and Rao

. (2014). Soliciting strategies for developing cell-based reference materials to advance mesenchymal stromal cell research and clinical translation. Stem Cells Dev, 23:1157–1167.

24.

Weiss

, Rao

, Deans

and Czermak

. (2016). Manufacturing cells for clinical use. Stem Cells Int, 2016:1750697.

25.

Bertolo

, Steffen

, Malonzo-Marty

and Stoyanov

. (2015). Canine mesenchymal stem cell potential and the importance of dog breed: Implication for cell-based therapies. Cell Transplant, 24:1969–1980.

26.

de Bakker

, Van Ryssen

, De Schauwer

and Meyer

. (2013). Canine mesenchymal stem cells: state of the art, perspectives as therapy for dogs and as a model for man. Vet Q, 33:225–233.

27.

Black

, Gaynor

, Gahring

, Adams

, Aron

, Harman

, Gingerich

and Harman

. (2007). Effect of adipose-derived mesenchymal stem and regenerative cells on lameness in dogs with chronic osteoarthritis of the coxofemoral joints: a randomized, double-blinded, multicenter, controlled trial. Vet Ther, 8:272–284.

28.

Guercio

, Di Bella

, Casella

, Di Marco

, Russo

and Piccione

. (2013). Canine mesenchymal stem cells (MSCs): characterization in relation to donor age and adipose tissue-harvesting site. Cell Biol Int, 37:789–798.

29.

Vilar

, Morales

, Santana

, Spinella

, Rubio

, Cuervo

, Cugat

and Carrillo

. (2013). Controlled, blinded force platform analysis of the effect of intraarticular injection of autologous adipose-derived mesenchymal stem cells associated to PRGF-Endoret in osteoarthritic dogs. BMC Vet Res, 9:131.

30.

Upchurch

, Renberg

, Roush

, Milliken

and Weiss

. (2016). Effects of administration of adipose-derived stromal vascular fraction and platelet-rich plasma to dogs with osteoarthritis of the hip joints. Am J Vet Res, 77:940–951.

31.

Cabon

, Febre

, Gomez

, Cachon

, Pillard

, Carozzo

, Saulnier

, Robert

, Livet

, et al. (2019). Long-term safety and efficacy of single or repeated intra-articular injection of allogeneic neonatal mesenchymal stromal cells for managing pain and lameness in moderate to severe canine osteoarthritis without anti-inflammatory pharmacological support: pilot clinical study. Front Vet Sci, 6:10.

32.

Seo

, Jeong

, Park

, Rho

, Kim

, Yu

, Lee

, Jung

, Lee

and Kang

. (2009). Isolation and characterization of canine umbilical cord blood-derived mesenchymal stem cells. J Vet Sci, 10:181–187.

33.

Sahoo

, Das

and Nayak

. (2017). Isolation, culture, characterization, and osteogenic differentiation of canine endometrial mesenchymal stem cell. Vet World, 10:1533–1541.

34.

Zucconi

, Vieira

, Bueno

, Secco

, Jazedje

, Ambrosio

, Passos-Bueno

, Miglino

and Zatz

. (2010). Mesenchymal stem cells derived from canine umbilical cord vein—a novel source for cell therapy studies. Stem Cells Dev, 19:395–402.

35.

Dominici

, Le Blanc

, Mueller

, Slaper-Cortenbach

, Marini

, Krause

, Deans

, Keating

, Prockop

and Horwitz

. (2006). Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy, 8:315–317.

36.

Escalhao

CCM

, Ramos

, Hochman-Mendez

, Brunswick

THK

, Souza

SAL

, Gutfilen

, Dos Santos Goldenberg

and Coelho-Sampaio

. (2017). Safety of allogeneic canine adipose tissue-derived mesenchymal stem cell Intraspinal transplantation in dogs with chronic spinal cord injury. Stem Cells Int, 2017:3053759.

37.

Taroni

, Cabon

, Febre

, Cachon

, Saulnier

, Carozzo

, Maddens

, Labadie

, Robert

and Viguier

. (2017). Evaluation of the effect of a single intra-articular injection of allogeneic neonatal mesenchymal stromal cells compared to oral non-steroidal anti-inflammatory treatment on the postoperative musculoskeletal status and gait of dogs over a 6-month period after tibial plateau leveling osteotomy: a pilot study. Front Vet Sci, 4:83.

38.

Eslaminejad

and Taghiyar

. (2010). Study of the structure of canine mesenchymal stem cell osteogenic culture. Anat Histol Embryol, 39:446–455.

39.

Choi

, Choi

, Kim

, Lee

, Park

, Kim

, Li

, Oh

, Lee

and Kim

. (2013). Isolation of canine mesenchymal stem cells from amniotic fluid and differentiation into hepatocyte-like cells. In Vitro Cell Dev Biol Anim, 49:42–51.

40.

Csaki

, Matis

, Mobasheri

, Ye

and Shakibaei

. (2007). Chondrogenesis, osteogenesis and adipogenesis of canine mesenchymal stem cells: a biochemical, morphological and ultrastructural study. Histochem Cell Biol, 128:507–520.

41.

Whitworth

, Frith

, Ovchinnikov

, Cooper-White

and Wolvetang

. (2014). Derivation of mesenchymal stromal cells from canine induced pluripotent stem cells by inhibition of the TGFbeta/activin signaling pathway. Stem Cells Dev, 23:3021–3033.

42.

Vieira

, Brandalise

, Zucconi

, Secco

, Strauss

and Zatz

. (2010). Isolation, characterization, and differentiation potential of canine adipose-derived stem cells. Cell Transplant, 19:279–289.

43.

Perez-Merino

, Uson-Casaus

, Zaragoza-Bayle

, Duque-Carrasco

, Marinas-Pardo

, Hermida-Prieto

, Barrera-Chacon

and Gualtieri

. (2015). Safety and efficacy of allogeneic adipose tissue-derived mesenchymal stem cells for treatment of dogs with inflammatory bowel disease: clinical and laboratory outcomes. Vet J, 206:385–390.

44.

Kisiel

, McDuffee

, Masaoud

, Bailey

, Esparza Gonzalez

and Nino-Fong

. (2012). Isolation, characterization, and in vitro proliferation of canine mesenchymal stem cells derived from bone marrow, adipose tissue, muscle, and periosteum. Am J Vet Res, 73:1305–1317.

45.

Takemitsu

, Zhao

, Yamamoto

, Harada

, Michishita

and Arai

. (2012). Comparison of bone marrow and adipose tissue-derived canine mesenchymal stem cells. BMC Vet Res, 8:150.

46.

Chow

, Johnson

, Coy

, Regan

and Dow

. (2017). Mechanisms of immune suppression utilized by canine adipose and bone marrow-derived mesenchymal stem cells. Stem Cells Dev, 26:374–389.

47.

Hong

, He

, Bui

, Ryba-White

, Rumi

, Soares

, Dutta

, Paul

, Kawamata

, et al. (2013). A focused microarray for screening rat embryonic stem cell lines. Stem Cells Dev, 22:431–443.

48.

Kobayashi

, Ichihara

, Sato

, Umeda

, Fields

, Fukumitsu

, Tago

, Ito

, Kainuma

, et al. (2019). On-site fabrication of Bi-layered adhesive mesenchymal stromal cell-dressings for the treatment of heart failure. Biomaterials, 209:41–53.

49.

Carrion

, Janson

, Kong

and Putnam

. (2014). A safe and efficient method to retrieve mesenchymal stem cells from three-dimensional fibrin gels. Tissue Eng Part C Methods, 20:252–263.

50.

Nawrocka

, Kornicka

, Szydlarska

and Marycz

. (2017). Basic fibroblast growth factor inhibits apoptosis and promotes proliferation of adipose-derived mesenchymal stromal cells isolated from patients with type 2 diabetes by reducing cellular oxidative stress. Oxid Med Cell Longev, 2017:3027109.

51.

Colenci

, da Silva Assuncao

, Mogami Bomfim

, de Assis Golim

, Deffune

and Penha Oliveira

. (2014). Bone marrow mesenchymal stem cells stimulated by bFGF up-regulated protein expression in comparison with periodontal fibroblasts in vitro. Arch Oral Biol, 59:268–276.

52.

Fierro

, Kalomoiris

, Sondergaard

and Nolta

. (2011). Effects on proliferation and differentiation of multipotent bone marrow stromal cells engineered to express growth factors for combined cell and gene therapy. Stem Cells, 29:1727–1737.

53.

Seshareddy

, Troyer

and Weiss

. (2008). Method to isolate mesenchymal-like cells from Wharton's Jelly of umbilical cord. Methods Cell Biol, 86:101–119.

54.

Devireddy

, Myers

, Screven

, Liu

and Boxer

. (2019). A serum-free medium formulation efficiently supports isolation and propagation of canine adipose-derived mesenchymal stem/stromal cells. PLoS One, 14:e0210250.

55.

Alves

, Serakides

, Boeloni

, Rosado

, Ocarino

, Oliveira

, Goes

and Rezende

. (2014). Comparison of the osteogenic potential of mesenchymal stem cells from the bone marrow and adipose tissue of young dogs. BMC Vet Res, 10:190.

56.

Ayala-Cuellar

, Kim

, Hwang

, Kang

, Lee

, Cho

and Choi

. (2019). Characterization of canine adipose tissue-derived mesenchymal stem cells immortalized by SV40-T retrovirus for therapeutic use. J Cell Physiol, 234:16630–16642.

57.

Bearden

, Huggins

, Cummings

, Smith

, Gregory

and Saunders

. (2017). In-vitro characterization of canine multipotent stromal cells isolated from synovium, bone marrow, and adipose tissue: a donor-matched comparative study. Stem Cell Res Ther, 8:218.

58.

Gardin

, Ferroni

, Bellin

, Rubini

, Barosio

and Zavan

. (2018). Therapeutic potential of autologous adipose-derived stem cells for the treatment of liver disease. Int J Mol Sci 19:pii:. E4064.

59.

Kang

, Kang

, Koo

, Park

, Choi

and Park

. (2008). Soluble factors-mediated immunomodulatory effects of canine adipose tissue-derived mesenchymal stem cells. Stem Cells Dev, 17:681–693.

60.

Liu

, Screven

, Boxer

, Myers

and Devireddy

. (2018). Characterization of canine adipose-derived mesenchymal stromal/stem cells in serum-free medium. Tissue Eng Part C Methods, 24:399–411.

61.

Mielcarek

, Storb

, Georges

, Golubev

, Nikitine

, Hwang

, Nash

and Torok-Storb

. (2011). Mesenchymal stromal cells fail to prevent acute graft-versus-host disease and graft rejection after dog leukocyte antigen-haploidentical bone marrow transplantation. Biol Blood Marrow Transplant, 17:214–225.

62.

Rodriguez Sanchez

, de Lima Resende

, Boff Araujo Pinto

, de Carvalho Bovolato

, Possebon

, Deffune

and Amorim

. (2019). Canine adipose-derived mesenchymal stromal cells enhance neuroregeneration in a rat model of sciatic nerve crush injury. Cell Transplant, 28:47–54.

63.

Ryu

, Lim

, Byeon

, Park

, Seo

, Lee

, Kim

, Kang

and Kweon

. (2009). Functional recovery and neural differentiation after transplantation of allogenic adipose-derived stem cells in a canine model of acute spinal cord injury. J Vet Sci, 10:273–284.

64.

Sasaki

, Mizuno

, Ozeki

, Katano

, Otabe

, Tsuji

, Koga

, Mochizuki

and Sekiya

. (2018). Canine mesenchymal stem cells from synovium have a higher chondrogenic potential than those from infrapatellar fat pad, adipose tissue, and bone marrow. PLoS One, 13:e0202922.

65.

Wood

, Chung

, Park

, Zwingenberger

, Reilly

, Ly

, Walker

, Vernau

, Hayashi

, et al. (2012). Periocular and intra-articular injection of canine adipose-derived mesenchymal stem cells: an in vivo imaging and migration study. J Ocul Pharmacol Ther, 28:307–317.

66.

Wang

, Yang

, Qi

and Jiang

. (2015). Osteoinduction and proliferation of bone-marrow stromal cells in three-dimensional poly (epsilon-caprolactone)/hydroxyapatite/collagen scaffolds. J Transl Med, 13:152.

67.

Tan

, Dinnes

, Myers

and Cooper-White

. (2011). Effects of biomimetic surfaces and oxygen tension on redifferentiation of passaged human fibrochondrocytes in 2D and 3D cultures. Biomaterials, 32:5600–5614.

68.

Spencer

and Lallier

. (2009). Mechanical tension alters semaphorin expression in the periodontium. J Periodontol, 80:1665–1673.

69.

Olsen

, Ng

, Lock

, Ahsan

and Rowley

. (2018). Peak MSC-are we there yet?. Front Med (Lausanne), 5:178.

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