Abstract
Human mesenchymal stem cells (hMSCs) promote endogenous tissue
regeneration and have become a promising candidate for cell therapy.
However, in vitro culture expansion of hMSCs induces a rapid decline of
stem cell properties through replicative senescence. Here, we
characterize metabolic profiles of hMSCs during expansion. We show that
alterations of cellular nicotinamide adenine dinucleotide (NAD + /NADH)
redox balance and activity of the Sirtuin (Sirt) family enzymes
regulate cellular senescence of hMSCs. Treatment with NAD + precursor
nicotinamide increases the intracellular NAD + level and re-balances
the NAD + /NADH ratio, with enhanced Sirt-1 activity in hMSCs at high
passage, partially restores mitochondrial fitness and rejuvenates
senescent hMSCs. By contrast, human fibroblasts exhibit limited
senescence as their cellular NAD + /NADH balance is comparatively
stable during expansion. These results indicate a potential metabolic
and redox connection to replicative senescence in adult stem cells and
identify NAD + as a metabolic regulator that distinguishes stem cells
from mature cells. This study also suggests potential strategies to
maintain cellular homeostasis of hMSCs in clinical applications.
Subject terms: Stem-cell biotechnology, Tissue engineering, Senescence
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Yuan et al. characterise metabolic profiles of human mesenchymal stem
cells (hMSCs) during cell expansion in culture. They find that late
passage hMSCs exhibit a NAD + /NADH redox cycle imbalance and that
adding NAD + precursor nicotinamide restores mitochondrial fitness and
cellular homeostasis in senescent hMSCs indicating a possible route to
preserve hMSC homeostasis for therapeutic use.
Introduction
In the past decades, human mesenchymal stem/stromal cells (hMSCs) have
become attractive candidates for cell therapy, as they exhibit
multi-lineage differentiation, paracrine effects, and
immunomodulation^[38]1,[39]2. For clinical applications, preparation of
hMSCs with translational and therapeutic standards represents a major
effort in cell therapy^[40]1,[41]3,[42]4. However, inconsistency of
clinical results of hMSCs has been observed, possibly due to donor
age/morbidity, isolation methods, and extensive in vitro culture
expansion^[43]5. For example, a negative phase III trial of random
donor MSCs in steroid-resistant graft-versus-host disease was
reported^[44]5,[45]6. hMSCs from aged or disease donors exhibit reduced
stemness and altered therapeutic efficacy including impaired paracrine
effects and immunomodulatory function^[46]7–[47]9. In addition, studies
have shown a culture-induced decline of stem cell properties in hMSCs
under prolonged culture^[48]1,[49]10. Extensive passaging of hMSCs
reduces the number of colony-forming unit-fibroblasts (CFU-F) and
proliferation rate^[50]10–[51]12, which is the hallmark of replicative
senescence^[52]13, corresponding with a gradual loss of therapeutic
potency in preclinical and clinical studies^[53]1. However, the
mechanism(s) that underpins the culture-induced senescence in hMSC has
not been well established.
Recent studies have demonstrated that hMSCs exhibit metabolic
plasticity both in vivo and in vitro, which contributes to cellular
properties and aging^[54]14,[55]15. hMSCs exhibit heterogeneity not
only at phenotypic level, but also at primary metabolic state
determined by their tissue origin^[56]15,[57]16. Upon isolation of
hMSCs from in vivo niche for expansion, in vitro nutrient-enriched
environment supports rapid cell proliferation, which requires energy
and anabolic macromolecules for daughter cell replication. In this
process, catabolic and anabolic pathways are interconnected and
together play essential roles in providing energetic sources as well as
metabolites to maintain cellular homeostasis^[58]17. Under this
context, hMSCs exhibit metabolic plasticity and can alter their
metabolic profile towards efficient oxidative phosphorylation (OXPHOS),
which is drastically different from quiescent glycolysis in their in
vivo niche^[59]18. Beyond energetic support, metabolic circuits engage
master genetic programs, and intermediate metabolites also mediate cell
signaling and regulatory pathways^[60]19. Thus, metabolic plasticity
allows hMSCs to match divergent demands for stem cell properties
including self-renewal and differentiation^[61]15,[62]16,[63]19. Our
previous studies indicated specific metabolic reconfigurations of hMSCs
in response to certain in vitro preconditioning conditions, which
further enhance stem cell functions. For instance, hMSCs under colonial
density or three-dimensional aggregation culture reconfigure the
energetic metabolism towards glycolysis, and thus improved hMSC
stemness^[64]16,[65]20. However, metabolic alterations associated with
replicative senescence in hMSCs during in vitro expansion have not been
well investigated.
As hMSCs utilize both glycolysis and OXPHOS as energy source to support
their proliferation and regenerative functions, intermediate
metabolites could regulate specific signaling pathways^[66]15,[67]16.
In particular, NAD^+/NADH redox cycle, which is integral to energy
production via glycolysis, tricarboxylic acid (TCA) cycle, and
OXPHOS^[68]21, is also a co-substrate of Sirtuin enzymes that regulate
cellular homeostasis and lifespan of organism, connecting central
energy metabolism to cellular aging and longevity^[69]21–[70]23.
Sirtuins (e.g., Sirt-1) utilize NAD^+ to catalyze the deacetylation of
histones in target proteins involved in aging process, including P53,
poly(ADP-ribose) polymerases (PARPs), the forkhead box O family
(FOXOs), nuclear factor-kB (NF-kB), peroxisome proliferator-activated
receptor-γ coactivator (PGC)-1α, and Ku70^[71]24,[72]25. Indeed, NAD^+
is the rate-limiting substrate in these Sirtuin pathways^[73]26. Change
to the intracellular NAD^+ level has been shown to influence cellular
metabolism and Sirtuins associated with aging^[74]27,[75]28. Other
signaling pathways in non-redox reactions also consume NAD^+ to
activate downstream functions, such as ADP-ribosyltransferases (e.g.,
PARPs) and cyclic ADP-ribose synthases (e.g., CD38, CD157), which
appear to be the major pathways to reduce the NAD^+
level^[76]21,[77]29. An increasing body of evidence has associated
decreasing intracellular NAD^+ level and Sirt-1 activity with metabolic
diseases and stem cell aging in vivo^[78]30. Strategies targeting the
activity of key enzymes involved in NAD metabolism, such as
nicotinamide phosphoribosyltransferase (NAMPT), CD38, and CD73,
demonstrate promising results in extending rodent healthspan or
lifespan^[79]21,[80]30,[81]31. Yet, the NAD^+/NADH redox balance along
with culture expansion, and the relations of NAD^+/NADH -Sirtuins and
replicative senescence in hMSCs remain to be understood.
Considering the evidence for hMSC metabolic plasticity and the role of
NAD^+/NADH in metabolism, this study tested the hypothesis that in
vitro culture expansion induces replicative senescence and metabolic
alterations in hMSCs which correlate with the loss of NAD^+ homeostasis
and result in the reduction of Sirt-1 signaling activity. In addition,
repletion of NAD^+ in senescent hMSCs recovers mitochondrial fitness
and glycolytic phenotype. Moreover, fully differentiated cell lines
such as human dermal fibroblasts (hFBs) do not exhibit culture-induced
senescence and changes in NAD^+ metabolism during culture expansion, as
hFBs and hMSCs are phenotypically similar^[82]32. Together, our study
reveals a novel metabolic and biochemical indicator which can be used
for hMSC quality control and rejuvenation in biomanufacturing.
Results
hMSCs become senescent and exhibit functional decline during in vitro
expansion
Since hMSCs are density-sensitive, consecutive passaging of hMSCs is
necessary to maintain them at the optimal density range (1000–6000
cells/cm^2). Our culture protocols for hMSC expansion followed the most
widely applied culture strategies (i.e., α-MEM plus 10% fetal bovine
serum, 80–90 confluence for harvest etc., details in Methods section).
In this study, hMSCs were found to exhibit significant morphological
changes and progressive alteration from small, spindle-shaped cells at
early passage (e.g., passage 5, referred as P5) to enlarged,
flat-shaped cells at late passage (e.g., passage 12, referred as P12)
(Fig. [83]1a). The enlarged cell size of hMSCs indicates increased
cellular senescence, which was also characterized by significantly
increased SA-β-gal activity (i.e., ~15% for P5 cells vs. 65% for P12
cells) in late passage of hMSCs (Fig. [84]1b, c). In addition,
increased DNA damage was observed using the Comet assay during culture
expansion, indicated by increased tail/body length in hMSCs at P12
compared to cells at P5 (Fig. [85]1d). hMSCs at late passage also
exhibited increased population doubling (PD) time (~6 days of P12 vs.
2.8 days of P5) (Fig. [86]1e), reduced capacity for self-renewal (~10
CFU-F colonies at P12 vs. 96 colonies at P5; Fig. [87]1f),
downregulation of stem cell genes Oct4 and Sox2 (but not Nanog)
compared to early passage cells (P5) (Fig. [88]1g). The increased
levels of mRNA expression for p53, p15, and p21 (Fig. [89]1h) and
accumulation of cells in G0/G1 phase of the cell cycle (Fig. [90]1i) in
P12 hMSCs, indicating a potential cell cycle arrest in hMSCs with
replicative senescence. Moreover, downregulation of autophagic genes,
including TFEB, BECN1, and LAMP1 were observed in hMSCs at P12
(Fig. [91]1j), corresponding to the significant decrease of autophagic
flux following culture expansion (Fig. [92]1k). Loss of autophagy
indicated the breakdown of cellular homeostasis of hMSCs during in
vitro culture expansion. For immunomodulation of hMSCs, mRNA expression
of genes (NF-kB and COX2) involved in chronic inflammation was found to
be upregulated in P12 hMSCs compared to P5 cells, while the
immunosuppressive ability via IDO pathways in P12 hMSCs decreased
(2.76-fold vs. 4.41-fold) under interferon-γ priming (Supplementary
Fig. [93]S1). The levels of potent anti-inflammatory cytokines HGF and
IL-10 decreased while the pro-inflammatory and anti-angiogenic CXCL10
cytokine level significantly increased in late passage of hMSCs. The
cytokine level of IL-6, TNF-α, and IL-1β was comparable for P5 and P12
cells (Supplementary Fig. [94]S2). Collectively, these results indicate
that hMSCs after prolonged culture expansion enter replicative
senescence, which disrupts cellular homeosasis and further reduces hMSC
function.
Fig. 1. In vitro culture expansion of human mesenchymal stem cells (hMSCs)
results in cellular senescence and breakdown of cellular homeostasis.
[95]Fig. 1
[96]Open in a new tab
a Alteration of hMSC morphology during culture expansion. b SA-β-gal
activity and c SA-β-gal staining in hMSCs at early passage and late
passage. d Comet assay demonstrates DNA damage of hMSCs during culture
expansion. e Population doubling (PD) time increased for long-term
cultured hMSCs. f Colony-forming unit-fibroblast (CFU-F) ability
decreased during culture expansion of hMSCs. g mRNA levels of stem cell
genes and h mRNA levels of cell cycle genes in hMSCs at late passage
compared to cells at early passage. i Cell cycle analysis of hMSCs via
flow cytometry. j Autophagic gene expression of hMSCs during culture
expansion. k Basal autophagic flux was reduced in hMSCs at late passage
compared to early passage. Late passage of hMSCs: passage 12 (P12),
early passage of hMSCs: passage 5 (P5). Biological replicates (n):
n = 3 for the tests. Scale bar: 100 µm. *p < 0.05; **p < 0.01;
***p < 0.001.
Culture expansion induces mitochondrial dysfunction in hMSCs
Consistent with the substantial changes in cell morphology,
mitochondrial morphology was also significantly altered during culture
expansion: from small-size fragmented morphology in P5 hMSCs towards
fused, elongated shape in P12 cells (Fig. [97]2a). This morphological
change was found to correspond to the increase of mitochondrial mass in
late passage hMSCs (P12) (Fig. [98]2b). However, tetramethylrhodamine
(TMRM) staining for mitochondrial transmembrane potential (MMP) was
found to decrease during culture expansion of hMSCs (Fig. [99]2c),
indicating a loss of membrane integrity and impaired electron transfer
ability. The depolarization of mitochondrial membrane was potentially
associated with the accumulation of both mitochondrial and total
cellular levels of reactive oxygen species (ROS) (Fig. [100]2d, e) and
decreased electron transport chain complex I (ETC-I) activity in hMSCs
of late passages (Fig. [101]2f). Corresponding to the loss of
autophagy, late passage hMSCs exhibited the decreased mitophagic flux
compared to early passage cells (Fig. [102]2g). Expression of genes
(e.g., MFN1, MFN2, FIS1, and DNM1L) involved in mitochondrial
fusion/fission dynamics was downregulated in late passage cells
(Fig. [103]2h), as well as genes involved in mitochondrial biogenesis
(e.g., NRF1, NRF2, SDHB, UQCRC1, and COX5B) (Fig. [104]2i). Together,
these data indicate that replicative senescence induced by long-term
culture expansion has substantial impacts on mitochondrial function and
biogenesis in hMSCs.
Fig. 2. Culture expansion of human mesenchymal stem cells (hMSCs) induces
mitochondrial dysfunction.
[105]Fig. 2
[106]Open in a new tab
a Mitochondrial morphology was altered during culture expansion of
hMSCs. b Increased mitochondrial mass and c loss of mitochondrial
membrane intensity were observed in hMSCs at late passage, both
determined by flow cytometry. d Mitochondrial reactive oxygen species
(ROS) and e total ROS were also increased during in vitro culture
expansion of hMSCs. f Electron transport chain complex I (ETC-I)
activity was reduced in late passage of hMSCs compared to cells at
early passage. g Culture expansion induced loss of hMSC mitophagy, as
well as h mitochondrial fusion and fission dynamics determined by mRNA
levels. i Genes involved in mitochondrial biogenesis were decreased
during culture expansion of hMSCs. Scale bar: 50 µm. Ng negative
control. Biological replicates (n): n = 3. *p < 0.05; **p < 0.01;
***p < 0.001.
Culture expansion induces metabolic reconfiguration in hMSCs
Since mitochondrial fitness is associated with energy metabolism and
metabolic plasticity of hMSCs during the adaption of culture
environment, the metabolic state of hMSCs at early and late passage was
characterized. Glycolytic ATP production was found to be significantly
reduced in hMSCs at P12 though slight increase of total ATP was also
observed (Fig. [107]3a). Multiple genes involved in glycolysis (i.e.,
PDK1, HK2, PKM2, LDHA, and G6PD) and the pentose phosphate pathway
(6PGD) were also downregulated in P12 cells (Fig. [108]3b). By
comparison, expression of the trans-aldolase and trans-ketolase enzymes
of the pentose phosphate pathway (TALDO1) was increased in P12 hMSCs.
Interestingly, the lactate production/glucose consumption ratio was
relatively stable (around 2.0) across the expansion of hMSCs from P4 to
P13 (Fig. [109]3c). However, gas chromatography–mass spectrometry
(GC–MS) analysis of hMSCs at different passages indicated that the
internal level of lactate increased in the P12 cells (Fig. [110]3d).
^13C-glucose tracer experiments showed that the amount of lactate
metabolized from glucose decreased, while the amount of citrate
increased in P12 hMSCs, suggesting an increased coupling of glycolysis
and TCA cycle metabolism as the labeled carbon of glucose has higher
enrichment in metabolites from TCA cycle (Fig. [111]3e). Furthermore,
the relative isotopic enhancement of individual carbons in citrate
(called isotopomers) did not show significant changes for M2 and M3
between P5 and P12 cells, while the M1, M4, and M6 isotopomers were
significantly altered, which may indicate other carbon source (i.e.,
glutamate and glutamine) instead of glycolysis may also participate in
the citrate metabolism (Fig. [112]3e, f). Whole cell metabolism was
also monitored in real time using a Seahorse flux analyzer. It was
found that both extracellular acidification rate (ECAR) and oxygen
consumption rate (OCR) values increased in P12 cells compared to P5
cells (Supplementary Fig. [113]S3). However, P12 hMSCs exhibited more
OXPHOS metabolic profiles after stressed by oligomycin and FCCP
(Carbonyl cyanide-4 (trifluoromethoxy) phenylhydrazone), indicating a
metabolic shift from glycolysis towards OXPHOS during in vitro culture
expansion of hMSCs (Fig. [114]3g, h).
Fig. 3. Culture expansion induces human mesenchymal stem cell (hMSC)
metabolic reconfiguration.
[115]Fig. 3
[116]Open in a new tab
a Glycolytic ATP ratio was decreased during culture expansion of hMSCs.
b mRNA level of genes in glycolysis and pentose phosphate pathways
(PPP) of hMSCs during culture expansion. c Lactate (Lac)
production/glucose (Glc) consumption ratio of hMSCs during extensive
expansion. Gas chromatography–mass spectrometry (GC–MS) analysis of
hMSCs during culture expansion. d Internal normalized peak area of
lactate. e Absolute total molar percent enrichment (ATMPE) levels of
^13C-glucose atoms in metabolites involved in glycolysis and OXPHOS. f
The relative molar percent enrichment (RMPE) levels of citrate. g
OCR/ECAR ratio changes with time of culture in hMSCs at early and late
passage. h hMSC energy metabolic phenotype during culture expansion
based on OCR and ECAR, open square: basal condition; closed square:
stressed condition. Biological replicates (n): n = 6 for GC–MS
metabolite study, n = 3 for rest of the tests. *p < 0.05; **p < 0.01.
To understand the global change of proteins involved in metabolism
during hMSC culture expansion, proteomic analysis of hMSCs at P4, P8,
and P12 was performed to illustrate the global difference in proteome
of hMSCs at early passage and late passage with replicative senescence
(Supplementary Fig. [117]S4–[118]S13). The experimental results
indicate 587 proteins in common (73–79%) across the P4, P8, and P12
hMSCs (Supplementary Fig. [119]S4A). Principal component analysis (PCA)
demonstrated a clear separation of P4, P8, and P12 samples as three
replicates of each group were clustered (Supplementary Fig. [120]S4B).
The 587 proteins were plotted in volcano plots and those with more than
10-fold change and those associated with NAD metabolism (e.g., HADHA,
NNMT, HSD17B4, OGDH, and DLD) were marked as yellow (Supplementary
Fig. [121]S4C). A total of 80 differentially expressed proteins (DEPs)
were selected based on 2-fold cutoff, p < 0.05 from the 587 proteins in
common. Systematic gene ontology (GO) enrichment analysis illustrated
the DEPs to be in three categories: (1) biological process category,
primarily enriched in metabolic process, biological regulation, and
response to stimulus; (2) cellular component category, primarily
enriched in membrane, membrane-enclosed lumen and nucleus; and (3)
molecular function category, primarily enriched in protein binding,
nucleic acid binding, and nucleotide binding (Supplementary
Fig. [122]S5). The DEPs were submitted to the Kyoto Encyclopedia of
Genes and Genomes (KEGG) database for intracellular pathway enrichment
analysis. A total of nine KEGG pathways were involved, in which the
metabolic pathways, glycolysis/gluconeogenesis, cysteine and methionine
metabolism, pyruvate metabolism, carbon metabolism, and PI3K-Akt
signaling pathway were mainly impacted (Supplementary Table [123]S1 and
Fig. [124]S6–[125]S8). These results are consistent with the
culture-induced metabolic reconfigurations observed using isotopic
tracer methods. Ingenuity Pathway Analysis (IPA) revealed top five
canonical pathways that are mostly impacted during culture expansion of
hMSCs, including EIF2 signaling, protein ubiquitination, fatty acid
beta-oxidation I, 2-ketoglutarate dehydrogenase complex, and superoxide
radicals degradation (Supplementary Fig. [126]S9). In particular, fatty
acid β-oxidation pathway and 2-ketoglutarate dehydrogenase complex
pathway are involved in energy metabolism and NAD^+/NADH redox cycle
(Supplementary Fig. [127]S10–[128]S13).
Culture expansion induces NAD^+/NADH redox cycle imbalance in hMSCs
The NAD^+/NADH redox cycle plays a crucial role in glycolysis and TCA
cycle and also participates in regulation of aging-related signaling
pathways and functions^[129]30. Since culture expansion of hMSCs
induces a metabolic reconfiguration of central metabolism, the redox
cycle balance may also be affected. Our results indicate that the
intracellular NAD^+ level progressively declined from P4, P9, to P12
cells, while the NADH level increased during in vitro culture expansion
(Fig. [130]4a). Since the NADH levels increased more than the NAD
levels decreased, ratio of NAD^+/NADH is lower in P12 hMSCs compared to
P4 cells (Fig. [131]4b). This culture-induced decline of NAD^+ level
and imbalance of NAD^+/NADH were further confirmed in multiple passages
and additional donors of hMSCs (Supplementary Fig. [132]S14). It was
also found that the protein level and gene expression of Sirt-1 and
Sirt-3, the key NAD^+-dependent enzymes that control cell signaling
pathways and function, were decreased in hMSCs at P12 (Fig. [133]4c, d,
e, f). Sirt-1 regulates mitochondrial biogenesis via PGC-1α and TFAM,
which were found to be downregulated in hMSCs at P12 compared to cells
at P5. In addition, the gene expression of PARP1, FOXO1, and FOXO3,
which are involved in oxidative stress and regulated by Sirt-1 and
Sirt-3, was increased in cells of P12 compared to cells at P5
(Fig. [134]4g, h). Western blot results confirm the decrease of Sirt-1,
Sirt-3, as well as PGC-1 from P5 to P12 at the protein level
(Fig. [135]4i). Together, these results indicate that culture expansion
leads to the progressive decline of intracellular NAD^+ level and the
increase of NADH level, which together change the redox cycle balance
in high passage of hMSCs. The reduced expression of NAD^+-dependent
Sirtuin enzymes was unable to regulate mitochondrial fitness, DNA
repair, and other aging-associated pathways during hMSC culture
expansion.
Fig. 4. NAD^+/NADH-Sirtuin (Sirt) axis imbalance induced by in vitro culture
expansion of human mesenchymal stem cells (hMSCs).
[136]Fig. 4
[137]Open in a new tab
a Intracellular NAD^+ and NADH levels were altered and b NAD^+/NADH
ratio decreased during culture expansion of hMSCs. c
Immunocytochemistry of Sirt-1 expression in culture-expanded hMSCs. d
Sirt-1 and e Sirt-3 protein levels characterized by flow cytometry. MFI
mean fluorescence intensity. f mRNA levels of Sirt-1 and Sirt-3 in
hMSCs. g Genes in DNA repair and mitochondria (PARP1, PGC1, and TFAM)
regulated by Sirt-1 in hMSCs at different passages. h FOXO pathways
(FOXO1 and FOXO3) regulated by Sirt-1 and Sirt-3, determined by RT-PCR.
i Sirt-1, Sirt-3, and PGC-1α protein levels during culture expansion of
hMSCs determined by Western blot. Biological replicates (n): n = 3.
*p < 0.05; **p < 0.01.
NAD^+ biogenesis and metabolism are altered during in vitro culture expansion
of hMSCs
To further investigate the changes of NAD^+ and NADH levels, several
proteins involved in major pathway of NAD^+ biosynthesis and
consumption in hMSCs were measured. NAMPT, the rate-limiting enzyme in
the NAD salvage pathway that is responsible for maintaining cellular
NAD^+ levels, was found to significantly increase in hMSCs of P12
compared to cells of P5-6, both at the mRNA level and at the protein
level (Fig. [138]5a, b). hMSCs at P12 also exhibited higher levels of
CD38 and CD73, the enzymes responsible for NAD^+ consumption, compared
to P5 cells (Fig. [139]5c, d). Western blot analysis confirmed the
increase of NAMPT, CD38, and CD73 in senescent hMSCs (Fig. [140]5e, f).
These results demonstrate that culture expansion alters the expression
of enzymes responsible for NAD metabolism, which may contribute to the
changes in the intracellular NAD^+ and NADH levels as well as their
ratio.
Fig. 5. In vitro culture expansion of human mesenchymal stem cells (hMSCs)
alters NAD^+ biogenesis and metabolism.
[141]Fig. 5
[142]Open in a new tab
a mRNA levels of NAD^+ metabolic enzymes NAMPT, CD38, and CD73 in hMSCs
at early and late passage. b NAMPT, c CD38, and d CD73 protein
expressions were all increased in late passage of hMSCs determined by
flow cytometry. NG negative control. e, f Western blot confirmed the
increase of NAMPT, CD38, and CD73 in late passage of hMSCs. Biological
replicates (n): n = 3. *p < 0.05. NS not statistically significant.
Re-balancing NAD^+/NADH redox cycle restores mitochondrial fitness and
cellular homeostasis in hMSCs with replicative senescence
Due to the observed relationship between NAD^+ metabolism and
aging-associated functions in hMSCs (as schematized in Supplementary
Fig. [143]S15), re-balancing NAD^+/NADH redox cycle may restore
mitochondrial fitness and cellular homeostasis, and rejuvenate hMSCs
with replicative senescence. Repletion of intracellular NAD^+ was
achieved by adding the NAD^+ precursor, nicotinamide (NAM), to the hMSC
culture media as our results demonstrate that there is no NAM in the
media (Supplementary Fig. [144]S16). The intracellular NAD^+ level and
the ratio of NAD^+/NADH significantly increased after 96 h of NAM
treatment (Fig. [145]6a, b). Sirt-1 and Sirt-3 expression was also
increased (Fig. [146]6c). More importantly, senescence was reduced as
indicated by reduced SA-β-gal activity in P12 hMSCs (Fig. [147]6d).
Correspondingly, colony-forming ability was recovered after short-term
NAM treatment (from 3 to 11 colonies) (Fig. [148]6e). After adding NAM,
the increase of cell population in the S phase of cell cycle was
observed (Supplementary Fig. [149]S17). Consequently, no significant
change of population doubling time was announced (data not shown). In
addition, the presence of NAM increased glycolytic ATP ratio in P12
cells (Fig. [150]6f), potentially indicating the increased activity of
glycolysis in central metabolism in hMSCs after NAM treatment.
Consistent with this, P12 hMSCs with NAM treatment was found to exhibit
increased basal autophagy level (Fig. [151]6g), as well as the
reduction of mitochondrial mass (Fig. [152]6h), re-polarized
mitochondrial membrane (increased TMRM staining in Fig. [153]6i), and
increased ETC-I activity (Fig. [154]6j). The mitophagy was also
improved (Fig. [155]6k) and the total ROS level decreased
(Fig. [156]6l) with NAM treatment in P12 hMSCs. Together, these results
indicate that replenishing intracellular NAD^+ level and maintaining
the NAD^+/NADH redox balance in senescent hMSCs increase Sirtuin
activity and thus restore mitochondrial fitness and autophagy/mitophagy
to maintain cellular homeostasis. These improvements lead to the
partial recovery from replicative senescence in hMSCs at high passage,
as well as the improved mitochondrial fitness that facilitates central
energy metabolism.
Fig. 6. Repletion of NAD^+ via NAM restores mitochondrial function and
preserves stem cell function in long-term cultured human mesenchymal stem
cells (hMSCs).
[157]Fig. 6
[158]Open in a new tab
Senescent hMSCs at late passages were treated with NAM for 96 h. a NAD
level was increased and NADH level was decreased. b NAD^+/NADH ratio
was increased. c Sirt-1 and Sirt-3 expressions were both increased. d
SA-β-gal activity was decreased. e Colony-forming ability (CFU-F) and f
glycolytic ATP ratio were also increased. g Basal autophagy was
restored in senescent hMSCs after NAM treatment. MFI mean fluorescence
intensity. Mitochondrial fitness was restored. h Mitochondrial mass was
decreased. i Mitochondrial transmembrane potential (MMP) and j electron
transport chain complex I (ETC-I) activity was increased. k Mitophagy
ability was restored. FCCP mitochondrial uncoupler carbonilcyanide
p-triflouromethoxyphenylhydrazone. l Total reactive oxygen species
(ROS) level was reduced as determined by flow cytometry. Biological
replicates (n): n = 3. *p < 0.05; **p < 0.01.
NAD^+/NADH redox cycle and mitochondrial fitness are relatively stable during
replicative expansion of human dermal fibroblasts
Since hMSCs exhibit significant changes induced by replicative
expansion, similar analysis was then performed for human dermal
fibroblasts (hFBs), which were chosen as a representative type of
mature adult cells. In contrast to hMSCs, extensive culture expansion
(up to passage 15) of hFBs showed no significant difference in
population doubling time (Fig. [159]7a) and SA-β-gal activity
(Fig. [160]7b), indicating that hFBs did not enter cell cycle arrest
and become senescent during replicative expansion. Cellular homeostasis
was also maintained because no change in autophagic flux was observed
(Fig. [161]7c). hFBs at P15 had similar levels of mitochondrial
activity and fitness compared to P4 cells, such as mitochondrial mass
(Fig. [162]7d), transmembrane potentials (Fig. [163]7e), and mitophagy
(Fig. [164]7f). More interestingly, intracellular NAD^+ level and
NAD^+/NADH ratio was relatively stable during culture expansion up to
15 passages (Fig. [165]7g). No significant changes were found in Sirt-1
and Sirt-3 expression during the expansion of hFBs as well
(Fig. [166]7h, i). Finally, the expression levels of NAMPT, CD38, and
CD73 were comparable throughout the expansion process (Fig. [167]7j),
indicating that NAD^+ biosynthesis and metabolism was well maintained
in hFBs during replicative expansion. Together, these data indicate
that hFBs do not exhibit replicative senescence and are able to
preserve cellular homeostasis in artificial culture environment.
Fig. 7. Human dermal fibroblasts (hFBs) under replicative expansion exhibit
limited cellular senescence, mitochondrial dysfunction, and NAD^+ decline.
[168]Fig. 7
[169]Open in a new tab
a Population doubling time of hFBs at early and late passage. b
SA-β-gal activity for culture-expanded hFBs. c No significant
difference for autophagic flux in P4 and P15 hFBs. Red line: negative
control. Orange line and blue line represents basal autophagy and with
autophagy inhibitor Bafliomycin-A (Baf-A), respectively. d
Mitochondrial mass and e mitochondrial transmembrane potential (MMP)
showed no difference for P4 and P15 hFBs. f Mitophagy also showed no
significant difference in hFBs during culture expansion. Red line:
negative control. Orange line and blue line represents untreated and
with mitochondrial uncoupler (FCCP) treatment, respectively. g
Intracellular NAD^+ and NADH levels, as well as NAD^+/NADH ratios are
relatively stable during culture expansion of hFBs. h Sirt-1 and i
Sirt-3 protein expression determined by flow cytometry. j The
expressions of NAD^+ metabolic enzymes NAMPT, CD38, and CD73 were all
comparable for hFBs at different passages as determined by flow
cytometry. Biological replicates (n): n = 3.
Discussion
Beyond multi-lineage differentiation, hMSCs exhibit paracrine and
immunomodulatory abilities that facilitate endogenous tissue
regeneration, thus are recognized as a potential therapeutic candidate.
For clinical purpose, in vitro large-scale expansion of hMSCs is a
necessary step to meet the requirement for cell number and dosages
while maintaining genetic stability and therapeutic efficacy^[170]1.
However, progressive loss of stem cell properties and genetic
alterations during in vitro culture of hMSCs have been widely
reported^[171]1,[172]10. This phenomenon is termed as replicative
senescence and is the major barrier for hMSCs to be an “off-the-shelf”
therapeutic product. This study provides full characterizations of
hMSCs during in vitro culture expansion and a novel mechanism
underlines replicative senescence, proposing a potential rejuvenation
strategy. Particularly, our study reveals the regulatory role of
metabolism and intermediate metabolites in hMSC aging and introduces
NAD^+/NADH redox balance to connect mitochondrial fitness with
replicative senescence in hMSCs.
In vitro culture conditions have been shown to significantly impact
cell properties. For hMSCs, a similar Hayflick limit was observed in
our studies and by other groups, with altered morphology and arrested
cell proliferation following extensive culture^[173]33–[174]37. In
fact, ultra-structure study of cellular organelles revealed endoplasmic
reticulum and matrix vesicles were also dysregulated after replicative
expansion^[175]38. Some reports describe a lower ability for
differentiation and decreased multipotency of senescent hMSCs. For
example, some studies showed increased osteogenesis and decreased
adipogenesis, while others showed well-preserved adipogenic potential
with diminished osteogenic differentiation^[176]33,[177]34,[178]39, as
observed in our study (Supplementary Fig. [179]S18). This may be due to
the different medium compositions but still revealed the disruption of
multipotency of hMSCs after extensive expansion. In our study, cell
cycle arrest in senescent hMSCs was announced by the gradual increase
of p53, p21, and p15 gene expression. Though DNA damage was considered
as the trigger of p53/p21 signaling pathway to initiate DNA repair,
independent activation of p53, p21, and p16 via autophagy and metabolic
regulation through AMP-activated protein kinase (AMPK) was also
proposed^[180]40–[181]42. Loss of autophagy has been attributed to
functional decline of aged stem cells: for example, aged muscle stem
cells and hematopoietic stem cells showed the impaired autophagy along
with loss of their regenerative potential, both in vitro and in
vivo^[182]43,[183]44. Restoring autophagy via rapamycin and spermidine
treatment partially restored stem cell functions in
vivo^[184]43,[185]44. An interesting phenomenon is that the
heterogeneity of aged stem cells also leads to differential autophagic
activity^[186]44, potentially explaining the reduced autophagy in hMSCs
during culture expansion as heterogeneity increases^[187]45. For the
first time, our results demonstrate a close link between gradual loss
of basal autophagy (and mitophagy), a hallmark of cellular
homeostasis^[188]46,[189]47, and the replicative senescence in hMSCs.
Our previous studies have extensively demonstrated the metabolic
plasticity of hMSCs under artificial culture
conditions^[190]15,[191]16,[192]48,[193]49. Upon removal from the in
vivo niche, hMSCs start to adapt to the in vitro environment by
utilizing both glycolysis and OXPHOS for ATP production. Cellular
homeostasis that contributes to pluripotency and clonal phenotype is
well maintained by the low level of glycolytic metabolism for stem
cells^[194]15,[195]16. As metabolism shifts from glycolysis towards
primarily OXPHOS, a breakdown of cellular homeostasis is expected due
to the accumulation of ROS and damaged organelles. Thus, the metabolic
state could also act as a hallmark of replicative senescence during
hMSC expansion. As shown in this study, both OCR and ECAR were
increased during culture expansion of hMSCs, contributing to the
slightly increased ATP production. However, the ratio of ATP generated
from glycolysis gradually decreased, indicating that hMSCs switch their
metabolism toward OXPHOS and other pathways to efficiently produce more
ATP in order to support extensive replication and maintain stem cell
properties^[196]45. This process may exhaust mitochondrial functions
and generate ROS and damaged organelles as autophagy is impaired in
senescent hMSCs^[197]50. Proteomics and metabolomics analysis in
current study also reveal the metabolic dysregulations in hMSCs
following culture expansion, such as the upregulated fatty acid
β-oxidation in late passage hMSCs. This observation may explain the
slight increase of ATP production during culture expansion of hMSCs,
with gradually decreased percentage of glycolytic ATP. Clearly, hMSCs
with extensive culture exhibit metabolic imbalance, which is a hallmark
of loss of homeostasis. Moreover, genomics analysis of hMSCs with
replicative senescence has demonstrated that genes involved in cell
differentiation and apoptosis are upregulated in senescent cells,
whereas genes involved in mitosis and proliferation are
downregulated^[198]51. Global omics analysis provides the evidence that
changes in replicative senescence may be a general property across hMSC
lines regardless of tissue source and culture conditions, but more
investigations of the universality of the hMSC replicative senescence
in response to extensive culture are still needed^[199]52,[200]53.
NAD^+ has been reported to be a regulatory intermediate metabolite
associated with aging in yeast and rodents, but few studies focus on
its role in in vitro culture of human cells and in vitro
senescence^[201]21,[202]54. Our results demonstrate a progressive
decline of intracellular NAD^+ level with extensive culture expansion
of hMSCs. As a redox cofactor, NAD plays a central role in energy
metabolism and also acts as a substrate for enzymes involved in
cellular signaling pathways, such as PARPs and Sirtuins^[203]55.
Recently, studies have connected cellular NAD^+ level to
aging-dependent cell functional decline in multiple
organs^[204]56,[205]57. Replenishment of intracellular NAD^+ level was
reported to significantly extend the lifespan of yeast, flies, worms,
and mice^[206]58–[207]62. It was reported that NAD^+ supplement
rejuvenates senescent muscle stem cells and neural stem cells in aged
mice via regulation of mitochondrial fitness and the unfolded protein
response to improve metabolic activity^[208]30. Interestingly, our
study did not observe significant change in AMPK activity, an energy
gauge for sensing energetic alteration. Thus, culture-induced
replicative senescence of hMSCs may differ from in vivo stem cell aging
in the context of energy production. For hMSCs, a metabolic shift
towards OXPHOS contributes to the changes in the NAD^+/NADH redox
balance as NAD^+ is rapidly converted to NADH by TCA cycle (by
isocitrate dehydrogenase, α-ketoglutarate dehydrogenase, and malate
dehydrogenase). Generally, NADH would be oxidized back to NAD^+ through
the electron transport chain^[209]63. However, our results show that
senescent hMSCs with damaged mitochondria exhibit depolarized
mitochondrial membrane and impaired ETC-I functions, which could
compromise the ability to metabolize NADH and maintain NAD^+/NADH redox
balance. This observation thus indicates that senescent hMSCs may shift
towards more reducing (or less oxidized) NAD^+/NADH cycle. Notably,
decreased ETC activity generally leads to decreased OCR, which is
opposite in our observations. This may be due to other cellular events
also regulating basal OCR, such as ATP turnover, proton leak, and
non-mitochondrial oxygen consumption (ROS formation)^[210]64. In
addition, loss of autophagy/mitophagy further creates a feedback loop
that facilitates the redox imbalance in our study. Accumulated DNA
damage and continuous activation of PARP during culture expansion could
also facilitate the NAD^+ depletion in senescence
hMSCs^[211]65–[212]68.
Two major mechanisms in aging-related NAD^+ decline have been proposed:
(1) decline of NAD^+ biosynthesis; and (2) increase of NAD^+
consumption by enzymes competing with each other for the cellular NAD^+
pool or chronological aging-induced enzymatic dysfunctions^[213]25. To
better understand the factors contributing to the metabolism of NAD^+
in senescent hMSCs, several key enzymes involved in NAD^+ biosynthesis
and consumption were investigated in this study. CD38 and CD73 were
highly upregulated, indicating the enhanced consumption of NAD^+ in
hMSCs with replicative senescence. Similar to the way that CD38 is
upregulated in immune cells under inflammatory environment, hMSC
senescence can be attributed to chorionic inflammation following in
vitro culture^[214]69,[215]70. To our surprise, NAMPT, a rate-limiting
intermediate enzyme in the major salvage pathway for NAD^+ biogenesis,
is also highly upregulated in hMSCs with replicative senescence. This
finding is seemingly contradictory to the decline of NAD^+ level but
can be explained by the absence of NAD^+ biosynthetic substrates (or
NAD precursors) such as NAM. Consistent with this explanation, NAM
supplement improves intracellular NAD^+ level and mitochondrial fitness
in senescent hMSCs, even with short-term treatment in our study. Based
on these results, the NAD^+/Sirt-1 axis dysfunction could be a
potential checkpoint for the loss of stemness and breakdown of
homeostasis in adult stem cells, and can be restored by supplying NAD^+
precursors for biosynthesis.
This study further examined whether human dermal fibroblasts exhibit
similar changes during culture expansion. Surprisingly, within a
similar number of population doublings, hFBs exhibit a relatively
consistent cell growth and β-gal activity, indicating that the cellular
senescence did not increase following the expansion of hFBs. Moreover,
NAD^+/NADH redox balance as well as Sirt-1/Sirt-3 activity was well
maintained in hFBs at late passage. Mitochondrial function and
autophagy/mitophagy were also comparable throughout culture expansion
in hFBs. These results demonstrate that hMSCs and hFBs have different
sensitivities to artificial culture environment under in vitro
expansion. Generally, fibroblasts were considered to share similar
phenotypic characteristics with MSCs^[216]32,[217]71,[218]72, including
lineage-specific differentiation and colony-forming ability, though
these properties are highly donor- and tissue
source-dependent^[219]71,[220]73. Studies have revealed that
fibroblasts can be cultured for 60–80 population doublings before
entering replicative senescence^[221]74, making them much more
replicative compared to hMSCs. Moreover, hFBs do not exhibit metabolic
reconfiguration under the nutrient-enriched culture environment. In
fact, switching to anaerobic metabolism mostly occurs in response to
serum starvation rather than reducing oxygen level in hFB
culture^[222]75. By comparison, hMSCs are extremely sensitive to their
culture environment including nutrients, oxidative stress, mechanical
stimuli, or even gravity^[223]76. hMSCs are able to adapt to different
culture environments and stimuli (e.g., hypoxia and cytokine
potentiation) to maintain stemness and cellular function. The adaption
process, in most cases, is required for engineering hMSCs with enhanced
therapeutic potentials^[224]77. In fact, this sensitivity provides the
possibility to engineer hMSCs with culture conditions instead of
genetic modification. For instance, hypoxia, 3D aggregation, and
cytokine priming can enhance hMSC properties for clinical purposes via
metabolic reconfiguration^[225]15,[226]49,[227]78–[228]80. hFBs,
however, may not be engineered by the metabolic preconditioning since
they are less sensitive to the artificial culture environment.
hMSCs under in vitro culture expansion exhibit replicative senescence,
accompanied with functional decline and loss of homeostasis, which is
regulated by a metabolic shift and a change of NAD^+/NADH redox
balance. In addition, mitochondrial damage and loss of autophagy also
contribute to the replicative senescence of hMSCs during expansion. Our
results demonstrate that repletion of NAD^+ via its precursor NAM
re-activates NAD^+/Sirt-1 axis to improve mitochondrial fitness and
further restores cellular homeostasis in hMSCs with replicative
senescence. This observation suggests a simple strategy for
manipulating culture conditions for biomanufacturing to maintain
desired therapeutic quality in hMSC-based therapy.
Methods
hMSC and hFB cultures
Frozen hMSCs from passage 0 to 2 were acquired from Tulane Center for
Gene Therapy. The hMSCs were isolated from the bone marrow of multiple
healthy donors with age 19–49 years old based on plastic adherence,
negative for CD3, CD14, CD31, CD45, and CD117 (all less than 2%) and
positive for CD73, CD90, CD105, and CD147 markers (all greater than
95%) and possess tri-lineage differentiation potential upon in vitro
induction^[229]48,[230]81. Informed consent was obtained from all
research participants. All experimental procedures and ethical
regulations have been reviewed and approved by Office for Human
Subjects Protection & Institutional Review Board in Florida State
University. hMSCs (1 × 10^6 cells/mL/vial) in freezing media containing
α-MEM, 2 mM L-glutamine, 30% fetal bovine serum (FBS), and 5% dimethyl
sulfoxide (DMSO) were thawed and cultured following the method
described in our prior publications^[231]16,[232]49,[233]80,[234]82.
Briefly, hMSCs were expanded and maintained in complete culture media
(CCM) containing α-MEM with 10% FBS (Atlanta Biologicals,
Lawrenceville, GA) and 1% Penicillin/Streptomycin (Life Technologies,
Carlsbad, CA) in a standard incubator at 37 °C with 5% CO[2] and 20%
O[2]. Culture medium was changed every three days. Cells were grown to
70–80% confluence and then harvested by incubation with 0.25%
trypsin/ethylenediaminetetraacetic acid (EDTA) (Invitrogen, Grand
Island, NY) at 37 °C for 4–7 min. Harvested cells were re-plated at a
density of 1500 cells/cm^2 and subcultured up to passage 15. For
comparison of cells at different passages, hMSCs from the same source
were used.
Primary human dermal fibroblasts (containing mitochondria,
PCS-201-012™) were purchased from American Type Culture Collection
(ATCC, Manassas, VA) and subcultured in CCM up to passage 15. All
reagents were purchased from Sigma Aldrich (St. Louis, MO) unless
otherwise noted.
Cell number, CFU-F, SA-β-Gal activity, comet assay, and glucose/lactate
measurements
Cell number was determined by Quant-iT™ PicoGreen kit (Invitrogen,
Grand Island, NY). Briefly, cells were harvested, lysed overnight using
proteinase K (VWR, Radnor, PA), and stained with Picogreen to allow
quantitation of cellular DNA. Fluorescence signals were measured using
a Fluror Count (PerkinElmer, Boston, MA). Population doubling time
(mean PD time) was determined through culture in each passage:
[MATH: MeanPDtime=t<
mrow>log2n
:MATH]
where t is culture time and n is the cell number fold increase during
culture time t.
For CFU-F assay, hMSCs were harvested and re-plated at the density of
15 cells/cm^2 on 60 cm^2 culture dish and cultured for another 14 days
in CCM. Cells were then stained with 20% crystal violet solution in
methanol for 15 min at room temperature (RT) and gently washed with
phosphate-buffered saline (PBS) for three times. The number of
individual colonies were counted manually. Cellular senescence was
evaluated by SA-β-Gal activity assay kit (Sigma, St. Louis, MO) as
described in manufacturer’s instructions.
Fresh and spent CCM were collected to determine glucose consumption and
lactate production by YSI 2500 Biochemistry Select Analyzer (YSI,Yellow
Spring, OH). Cellular DNA damage was measured by comet assay (Cell
Biolabs, Inc. San Diego, CA), following manufacturer’s instructions.
Mitochondrial morphology, mass and membrane potential, and ROS level
measurements
For mitochondrial morphology, hMSCs were incubated with 100 nM
MitoTracker Red CMXRos (Molecular Probe, Eugene, OR) in CCM at 37 °C
for 30 min. After washing with PBS, cells were fixed with 3.7%
formaldehyde at 37 °C for 15 min and then imaged with Olympus IX70
microscope.
For mitochondrial mass and MMP measurement, trypsinized hMSCs were
washed in warm Hank’s Balanced Salt Solution (HBSS). Cell suspension
was incubated with MitoTracker green FM or tetramethylrhodamine, methyl
ester (Molecular Probe, Eugene, OR) at 37 °C for mass and MMP staining,
respectively. Cells were then washed with HBSS and analyzed by flow
cytometry (BD Biosciences, San Jose, CA).
For ROS measurement, cell suspension was incubated with 25 µM
carboxy-H2DCFDA (Molecular Probe) at 37 °C for 30 min and total ROS was
determined using flow cytometry. For mitochondrial ROS measurement,
cell suspension was incubated with 5 µM MitoSOX Red (Molecular Probe)
at 37 °C for 10 min and analyzed using flow cytometry.
Immunocytochemistry, cell cycle, autophagy, and mitophagy measurements
Cells were harvested with 0.25% trypsin-EDTA solution, washed in PBS,
and then fixed at 4% paraformaldehyde (PFA) at RT for 15 min. Cells
were then permeabilized in 0.2% triton X-100 for 10 min at RT.
Non-specific binding sites were blocked with 1% bovine serum albumin,
10% FBS in PBS for 15 min at RT. After washing, cells were incubated
with specific primary antibodies for human Sirt-1, Sirt-3, NAMPT, CD38,
and CD73 (Santa Cruz Biotechnology, Dallas, TX) at RT for 2 h, followed
by incubation with FITC-conjugated secondary antibody (Molecular
Probe). Labeled samples were analyzed by flow cytometry. Antibody
information was summarized in Supplementary Table [235]S2. Gating
strategy for hMSCs at early and late passage was demonstrated in
Supplementary Fig. [236]S19.
For cell cycle analysis, suspended cells were fixed with 70% cold
ethanol for 30 min at 4 °C and then washed with PBS. 100 µg/mL RNase A
(VWR, Radnor, PA) was added to cell suspension and incubated at 35 °C
for 15 min. Then the samples were incubated with 400 µL 50 µg/mL of
propidium iodide (VWR) solution at RT in the dark for 1 h. Cell cycle
was then determined by flow cytometry.
For autophagy measurement, cell suspension was incubated with 20 µM
Cyto-ID Green (Enzo Life Sciences, Farmingdale, NY), a fluorescent dye
that selectively labels accumulated autophagic vacuoles, at 37 °C for
30 min, and analyzed by flow cytometry and calculated according to the
manufacturer’s instructions. For mitophagy measurement, the cells were
incubated with mitochondrial uncoupler carbonilcyanide
p-triflouromethoxyphenylhydrazone (FCCP, 1 µM) for 20 min and then the
mitochondrial mass was tested via flow cytometry. Mitophagic flux in
the cells was calculated by the different mitochondrial mass between
treated and untreated group.
Intracellular ATP content, mitochondrial complex I activity, and metabolic
phenotype
hMSCs were centrifuged, re-suspended in deionized water, and heated
immediately in boiling water for 15 min. The mixture was centrifuged,
and ATP-containing supernatant was collected. Upon measurement, 10 µL
of ATP solution was mixed with 100 µL of the luciferin-luciferase
reagent (Sigma-Aldrich), and the bioluminescent signal was measured
using an Orion Microplate Luminometer (Bad Wildbad, Deutschland). To
determine the glycolytic ATP ratio, cells were cultured with or without
glycolysis inhibitor 2-Deoxy-D-glucose (2-DG, 5 mM) for 48 h and then
the ATP was measured. The ratio of glycolytic ATP was calculated by the
delta value of total ATP and 2-DG treated ATP normalized to total ATP.
Activity of mitochondrial electron transport complex I activity was
determined using the Complex I Enzyme Activity Microplate Assay Kit
(Abcam, Cambridge, MA) according to the manufacturer’s instructions.
OCR and ECAR were determined using Agilent Seahorse XF Extracellular
Flux Analyzer XFp (Seahorse Biosciences, Massachusetts, USA). All tests
were performed in accordance with manufacturer’s instructions. Briefly,
hMSCs were seeded onto Seahorse XFp Cell Culture Miniplate (Seahorse
Biosciences) at 10,000 cells per well the day before being analyzed.
Cells were equilibrated in a non-CO[2] incubator with Seahorse
calibrant buffer for 60 min prior to assay. Using the Seahorse XFp Cell
Energy Phenotype Test Kit (Seahorse Biosciences), OCR and ECAR under
baseline and stressed conditions (oligomycin and FCCP) were
measured^[237]83.
Intracellular NAD^+ and NADH quantification
Intracellular NAD^+ and NADH were measured with NAD^+/NADH
Quantification Colorimetric Kit (BioVision, Milpitas, CA) according to
manufacturer’s instructions with some modifications. Briefly,
approximate 0.8 million cells were collected and directly lysed in
200 µL lysis buffer from the assay kit. The volume of reagents in each
step was scaling down by 50% and the results were calculated by the
freshly prepared standard curve (NADH standards provided by the assay
kit). Final NAD^+ and NADH concentrations were then normalized to the
total cell number in each group.
^13C-glucose labeling and GC–MS analysis of hMSC metabolites
^13C-glucose labeling, metabolite extraction, and chemical
derivatization were performed as in our prior
publications^[238]16,[239]48. Briefly, glucose-free DMEM medium
supplemented with a 2:3 mixture of unlabeled and U-^13C- labeled
glucose (Cambridge Isotopes Laboratories, Andover, MA) at the same
concentration as CCM for hMSC expansion (1.0 g/L glucose). P5 and P12
hMSCs were seeded and cultured for 2 days in DMEM with unlabeled
medium. The culture medium was then replaced with isotope-enriched
medium and cultured for additional 3 days. Cell collection started by
washing with PBS, quenching with liquid nitrogen, and addition of a
solution of methanol:water (4:1) directly to the culture plate to stop
metabolism and lyse the cells on dry ice, followed by addition of the
internal standard (norleucine 28 µg/ml solution) and incubation at
−80 °C for 10 min. The extracts were then centrifuged at 5000 x g for
5 min at 4 °C, and the supernatants were collected and transferred to a
silanized Reacti-Vial (Wheaton) and stored at −80 °C. Prior to
derivatization, frozen extracts were dried under vacuum overnight and
dissolved in 20 µL pyridine and 20 µL
N-methyl-N-(tert-butyldimethylsilyl) trifluoro-acetamide containing 1%
tert-butyldimethylchlorosilane (Thermo Scientific, Rockford, IL). The
reaction was performed under a stream of argon. Reacti-vials were
closed under argon and heated to 75 °C for 60 min and then cooled to
room temperature. Injection of derivatized extracts in the GC–MS was
completed within 24 h of derivatization.
Derivatized samples (1 μL) were injected in splitless mode at 230 °C in
an HP Agilent 6890 series gas chromatograph (GC) coupled with an HP
Agilent 5973 mass selective detector and separated on a 30 m DB5 column
(J&W Scientific, Folsom, CA). The GC oven temperature was held at 70 °C
for 1 min after injection, increased to 120 °C at 15 °C min^−1 and
finally to 325 °C at 10 °C min^−1. Mass spectra were collected over m/z
50–650 at a rate of 2 Hz with MS source set at 260 °C. Metabolites were
identified by comparison with standards. Peak areas were calculated
from the [M-57]+• and [M-159]+• ions for amino acids and [M-57]+• and
[M-189]+• ions for carboxylic acids by fitting the elution profile of
each isotopomer to a Gaussian, eliminating the baseline, and summing
over all isotope peaks for each specific ion^[240]84,[241]85. The area
was then normalized to the peak area of the internal standard
norleucine, and divided by the cell count. Mass isotope distribution
vectors and isotope incorporation was determined using methods
described in detail elsewhere^[242]48.
Real-time reverse transcriptase-polymerase chain reactions
Total RNA was isolated using the RNeasy Plus kit (Qiagen) following
vendor’s instructions. Reverse transcription was carried out using 2 μg
of total RNA, anchored oligo-dT primers (Operon), and Superscript III
(Invitrogen). Primers for specific target genes were designed using the
software Oligo Explorer 1.2 (Genelink) (Supplementary Table [243]S3).
β-actin was used as an endogenous control for normalization. RT-PCR
reactions were performed on an ABI7500 instrument (Applied Biosystems),
using SYBR Green PCR Master Mix. The amplification reactions were
performed and the quality and primer specificity were verified. Fold
variations in gene expressions were quantified using the comparative Ct
method: 2^-Δ(CtTreatment -CtControl), which is based on the comparison
of the target gene (normalized to β-actin) among different conditions.
Statistics and reproducibility
Unless otherwise noted, all experiments were performed at least in
triplicate (n = 3), and representative data are reported. Experimental
results are expressed as means ± standard deviation (SD) of the
samples. Statistical comparisons were performed by one-way ANOVA and
Tukey’s post hoc test for multiple comparisons, and significance was
accepted at p < 0.05.
Supplementary information
[244]Peer Review File^ (449.7KB, pdf)
[245]Supplementary Information^ (2.5MB, pdf)
Acknowledgements