58-0459-41

antibody from Invitrogen Antibodies

Targeting: PTPRC CD45, GP180, LCA, T200

Provider product page for 58-0459-41

Immunohistochemistry

Flow cytometry

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Antibody data

Antibody Data
Antigen structure
References [40]
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Validations
Immunohistochemistry [2]
Flow cytometry [1]
Other assay [51]

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Product number: 58-0459-41 - Provider product page
Provider: Invitrogen Antibodies
Product name: CD45 Monoclonal Antibody (HI30), Alexa Fluor™ 532, eBioscience™
Antibody type: Monoclonal
Antigen: Other
Description: Description: The HI30 monoclonal antibody reacts with all isoforms of human CD45, also known as Leukocyte Common Antigen (LCA). CD45 is expressed by all hematopoietic cells excluding circulating erythrocytes and platelets. The cytoplasmic portion of CD45 has tyrosine phosphatase enzymatic activity and plays an important role in activation of lymphocytes. Applications Reported: This HI30 antibody has been reported for use in flow cytometric analysis. Applications Tested: This HI30 antibody has been pre-titrated and tested by flow cytometric analysis of normal human peripheral blood cells. This can be used at 5 µL (0.06 µg) per test. A test is defined as the amount (µg) of antibody that will stain a cell sample in a final volume of 100 µL. Cell number should be determined empirically but can range from 10^5 to 10^8 cells/test. Alexa Fluor® 532 is excited with the Green laser (532 nm) and emits at 561 nm. This cannot be used with the Yellow-Green laser (561 nm). We recommend using a 560/14 band pass filter. Please make sure that your instrument is capable of detecting this fluorochome. Excitation: 532 nm; Emission: 561 nm; Laser: Green Laser
Reactivity: Human
Host: Mouse
Conjugate: Yellow dye
Isotype: IgG
Antibody clone number: HI30
Vial size: 25 Tests
Concentration: 5 µL/Test
Storage: 4° C, store in dark, DO NOT FREEZE!

PTPRC protein structure - 58-0459-41 shown in red.

Supportive validation

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Experimental details: Immunohistochemical analysis of CD45 was performed using formalin-fixed paraffin-embedded human tonsil tissue sections. To expose the target protein, heat-induced epitope retrieval was performed on de-paraffinized sections using eBioscience™ IHC Antigen Retrieval Solution - High pH (10X) (Product # 00-4956-58) diluted to 1X solution in water in a decloaking chamber at 110 degree Celsius for 15 minutes. Following antigen retrieval, the sections were blocked with 2% normal goat serum in 1X PBS for 45 minutes at room temperature and then probed with or without CD45 Monoclonal Antibody (HI30), Alexa Fluor™ 532, eBioscience™ (Product # 58-0459-42) at 1:100 dilution in 0.1% normal goat serum overnight at 4 degree Celsius in a humidified chamber. ReadyProbes™ Tissue Autofluorescence Quenching Kit (Product # R37630) was used to quench autofluorescence from the tissues. Nuclei were stained with DAPI (Product # D1306) and the sections were mounted using ProLong™ Glass Antifade Mountant (Product # P36984). The images were captured on EVOS™ M7000 Imaging System (Product # AMF7000) at 20X magnification and externally deconvoluted.
Conjugate: Yellow dye

Supportive validation

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Experimental details: Immunohistochemical analysis of CD45 was performed using formalin-fixed paraffin-embedded human tonsil tissue sections. To expose the target protein, heat-induced epitope retrieval was performed on de-paraffinized sections using eBioscience™ IHC Antigen Retrieval Solution - High pH (10X) (Product # 00-4956-58) diluted to 1X solution in water in a decloaking chamber at 110 degree Celsius for 15 minutes. Following antigen retrieval, the sections were blocked with 2% normal goat serum in 1X PBS for 45 minutes at room temperature and then probed with or without CD45 Monoclonal Antibody (HI30), Alexa Fluor™ 532, eBioscience™ (Product # 58-0459-42) at 1:100 dilution in 0.1% normal goat serum overnight at 4 degree Celsius in a humidified chamber. ReadyProbes™ Tissue Autofluorescence Quenching Kit (Product # R37630) was used to quench autofluorescence from the tissues. Nuclei were stained with DAPI (Product # D1306) and the sections were mounted using ProLong™ Glass Antifade Mountant (Product # P36984). The images were captured on EVOS™ M7000 Imaging System (Product # AMF7000) at 20X magnification and externally deconvoluted.
Conjugate: Yellow dye

Supportive validation

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Experimental details: NULL
Conjugate: Yellow dye

Supportive validation

Submitted by: Invitrogen Antibodies (provider)
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Supportive validation

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Experimental details: NULL
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Supportive validation

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Experimental details: NULL
Conjugate: Yellow dye

Supportive validation

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Experimental details: NULL
Conjugate: Yellow dye

Supportive validation

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Experimental details: Figure 1 Inflammatory leukocyte infiltrates are present in rejected human allografts . (A) Schematic diagram illustrating our working model. In response to ischemic kidney injury or inflammation upon allograft transplantation, CD11b+ leukocytes are recruited into the tissue, where they induce an adaptive response and further recruitment of innate and adaptive immune cells. Increased inflammation results in progressive tissue damage, nephropathy and, eventually, graft failure. Given that CD11b+ leukocyte infiltration into the allograft is an early event, blocking influx of these innate immune cells via novel small molecule CD11b agonists is expected to reduce inflammation, decrease progressive tissue damage and prolong allograft survival. (B) Representative micrographs of human kidney tissue sections (from a healthy cadaveric donor or a rejected allograft) after histochemical and immunofluorescence analyses. Sequential sections from each tissue were stained with periodic acid-Schiff (PAS) or trichrome and imaged using light microscopy. Sections were also stained with antibodies specific for CD45, CD11b, or CD3, as indicated, followed by fluorescently labeled secondary antibodies and then imaged using a confocal microscope. The rejected allograft shows enhanced trichrome staining (indicating an increase in interstitial fibrosis) and increased numbers of CD45+, CD11b+, and CD3+ immune cells as compared to the healthy kidney ( n = 4-5/group). Scale bar = 50 mum.
Conjugate: Yellow dye

Supportive validation

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Experimental details: Figure 5 LA1 treatment reduces perivascular infiltration of macrophages . (A) Representative micrographs of kidney allograft tissue sections from control (CsA alone) and LA1-treated (CsA + LA1) animals after immunofluorescence analyses. Sequential sections from each tissue were stained with antibodies against CD45, CD11b, or F4/80, as indicated, followed by fluorescently labeled secondary antibodies and then imaged using a confocal microscope. Inset shows image of an area of 60x magnification. Images are representative of 4-5 independent samples per group. Scale bar = 50 mum. (B) Plots showing quantification of number of positive cells/area for each marker, as shown in (A) . Three random areas per tissue were evaluated at 60x magnification by manually counting positively stained cells. Data shown are mean +- SEM ( n = 4-5/group). Significance was determined using a two-tailed Student's t -test. *** P < 0.001; **** P < 0.0001.
Conjugate: Yellow dye

Supportive validation

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Experimental details: Fig. 10 Immunocytochemical analysis of CTCs that were isolated from the samples using Pancytokeratin magnetic beads after 48 and 72 h. DAPI was used to stain the cells' nucleus. Anti-CD45-PE antibody (orange) was used to stain PBMCs and anti pancytokeratin antibody to stain CTCs
Conjugate: Yellow dye

Supportive validation

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Experimental details: Fig. 11 Immunocytochemical analysis of CTCs that were isolated from the samples using pancytokeratin magnetic beads after 96 h. DAPI was used to stain the cells' nucleus. Anti-CD45-PE antibody (orange) was used to stain PBMCs and anti pancytokeratin antibody to stain CTCs
Conjugate: Yellow dye

Supportive validation

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Experimental details: Fig. 9 Immunocytochemical analysis of CTCs that were isolated from the samples using Pancytokeratin magnetic beads after 0 and 24 h. DAPI was used to stain the cells' nucleus. Anti-CD45-PE antibody (orange) was used to stain PBMCs and anti pancytokeratin antibody to stain CTCs
Conjugate: Yellow dye

Supportive validation

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Experimental details: Figure 1 Retinoic Acid Signaling Inhibition Increases the Yield of Hematopoietic Progenitors with an HSC-like Phenotype from Human Pluripotent Stem Cells (A) Schematic of pluripotent stem cell differentiation system used to model human hematopoietic development through mesoderm specification and blood commitment. RA inhibitors or RA was present continuously during the 16-day differentiation, except where otherwise stated. (B) Representative FACS plots showing the hematopoietic population derived from pluripotent stem cells at day 16 of differentiation. FACS gates show blood (CD45/43 + ), hematopoietic progenitors (CD45/43 + CD34 + ), and HSC-like immature progenitors (CD45/43 + CD34 hi CD38 lo CD90 + CD45RA - ). Gates are based on FMO controls with more stringent CD34 hi and CD38 lo gating based on cord blood hematopoietic stem and progenitor cell standards. Doublet exclusion and dead cell exclusion were done before applying the gates. (C) Frequency (%) of blood subfractions derived from cells treated with DEAB (green) at 10 muM from seven independent experiments (n = 7) (see also Figures S1 A and S1B) or RA (orange) (0.01, 0.1, 1.0 muM; n = 3). All values are normalized to the DMSO control (gray) as indicated by the dotted line (100%). X = no events. Bar graphs show change in cell viability as measured by 7AAD for tested conditions. (D) mRNA expression level of RA target gene RARbeta at day 8 of differentiation (n = 3). (E) Frequency of the HSC-like fraction in cells treated
Conjugate: Yellow dye

Supportive validation

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Experimental details: Figure 2. Feasible method to detect and isolate CTCs. (A) Gates were set to capture viable CTCs via FACS analysis. (A-b) The P1 gate was set to obtain cells with diameters larger than lymphocytes, and (A-c) the P2 gate was set to acquire granulocyte-like side scatter (SSC) cells. (A-a) P1 was combined with P2 to capture viable cells similar to tumour cells in size and SSC. (A-d and -f) The P3 and P4 gates were used to select hTERT-positive cells without leucocyte-common antigen (CD45) expression and collect highly hTERT-positive cells for further analysis. (A-e and -f) The sorted cells were detected at P5 gate (taking partial results from patient B1 for example). (A-i) The single-stained CD45-APC positive control of PBMC and (A-j) the infected control, including negative control (peripheral blood of a healthy person), (A-h) positive control (the hTERT-expressing SK-BR-3 cancer cell line). (B) Analysis of circulating breast tumour cells using imaging flow cytometry. (B-a) Cell image mapped with the point in the bottom right corner of R1 gate (high signal captured in ch02 and low signal captured in ch07, corresponding to the gate P5). Ch02 received signal emitted by GFP (hTERT), ch07 reflects the signal intensity of eFluor450 (CD45), ch11 represents the APC (ErbB2) intensity and ch12 is the SSC channel. (B-b) The merged image represents hTERT + CD45-ErbB2 + cells. (B-c) The copy number profiles of a near-triploid CTC.
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Supportive validation

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Experimental details: Figure 2 hWJMSC-MVs surface expressed molecules analysis. Flow cytometery analysis showed hWJMSC-MVs were positive for some surface expressed molecules typically expressed by MSCs, such as CD9, CD44, CD63, CD73, and negative for CD34, CD45.
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Supportive validation

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Experimental details: Extended Data Figure 6 Conditional expression of FGRS is sufficient for optimal generation of rEC-hMPPs with multilineage potential, including T-Cell lymphoid cells A to C: Conditional expression of mouse inducible FGRS factors activates endogenous human FGRS in HUVECs sustaining functional hematopoietic cell fate of rEC-hMPPs. A . To test whether FGRS-induced reprogramming triggered expression of endogenous FGRS genes , HUVECs were transduced with lentivirus expressing FGRS-Tet-On and a trans-activator, and grown on E4EC-vascular niche for 18-22 days (n=4) in the presence of doxycycline. Doxycycline was removed from the culture medium after 18-22 days to shut off the expression of mouse FGRS and cells were cultured for additional 7-10 days. Human CD45 + CD34 + cells were FACS isolated for CFC assay and whole-transcriptome deep sequencing (RNA-seq). CFC assay revealed emergence of hematopoietic colonies with cells expressing human CD235, CD11b, CD83, and CD14. B . Comparison of transcriptional profiles of the human FGRS expression in human HUVECs, hCD45 + rEC-hMPPs programmed using inducible mouse FGRS, CD45 + CD34 + rEC-hMPPs, 22 weeks post-transplantation, hDMEC-derived CD45 + CD34 + rEC - hMPPs after 15 weeks post-secondary engraftment and naive CD34 + Lin + cells purified from cord blood. C . Analysis of whole-transcriptome RNA-Seq of rEC-hMPPs derived using inducible mouse FGRS (n=3). All RNA-Seq reads were aligned against human and mouse FGRS sequences. RNA-Seq reads th
Conjugate: Yellow dye

Supportive validation

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Experimental details: Extended Data Figure 2 FGRS-transduction and vascular-induction reprogram HUVECs, but not hES-ECs, to proliferating functional rEC-MPPs A. Multi-colony niche-like structure that physically separates developing hematopoietic colonies from surrounding E4EC vascular niche . The emerging multi-colony sinusoidal-like structures create a unique cellular interface between E4EC monolayers and transduced ECs giving rise to hematopoietic clusters (n=4, scale bar is 1000 um.) B. Expansion potential of reprogrammed CD45 + hematopoietic cells . CD45 + (12x103) and CD45- (60 x10 3 ) cells were sorted into separate wells and expanded for two days. We observed 5-fold expansion of CD45 + cells (56.6 x10 3 +-7.9 x10 3 ; n=3) and dramatic reduction of CD45 - cells number (4.6x10 3 +-1.0 x10 3 ; n=3). C . Clonal expansion of CD45 + cells. CD45 + cells were FACS sorted into 96-well plates at the density of 1 and 2 cells/well. After seven days of culture, we observed CD45 + cell expansion in 6.3+-2.1 wells (93.1+-14.5 cells/well) of 1-cell sort and 29.0+-4.3 wells (112.1+-21.2 cells/well) of the 2-cell sort (n=3). The difference between cell number in 1 and 2-cell sort was statistically not significant (p=0.78) suggesting that the difference in the number of wells with detected cell expansion was rather due to survival of sorted cells than a reflection of the number of cells sorted into a well. D. FGRS-induced generation of hematopoietic cells by hES-derived endothelium (hES-EC) . Representative e
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Supportive validation

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Experimental details: Figure 3 Freeze/thaw of standard plasma increased platelet MPs in platelet poor plasma (PPP). Standard plasma from six healthy volunteers was either immediately processed to PPP or exposed to a single freeze/thaw cycle prior to a second centrifugation step to remove residual platelets. Platelet (CD41+), leukocyte (CD45+), endothelial (CD146+), and RBC (CD235a) MPs were quantified by flow cytometry. Total number of MPs in a sample was either calculated by the sum of positive events detected by antibody staining (Sum antibody+) or measured by staining with calcein-AM, a non-specific dye that fluorescently labels intact vesicles. Results were similar and showed marked increase after freeze-thaw of standard plasma *** indicates P < 0.001. This effect was largely due to an increase in platelet derived MPs (CD41+).
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Supportive validation

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Experimental details: Figure 2 T-cell-specific deletion of RIP1. ( a ) Genotyping by PCR of mouse ear DNA showing genotypes generated from a RIP1 +/f x RIP1 +/f Lckcre + mating cross. ( b ) Western blotting analysis of RIP1 in thymocytes from mice of the indicated genotypes. Ponceus S staining was performed as protein loading/transfer control. ( c ) Total organ cellularity of RIP1 t-/- mutant mice (filled circles) and RIP1 +/+ control mice (open circles) are shown. Error bars are average+-S.E.M. P -values >0.05. ( d ) Representative two-color flow cytometric plots showing the T cell (CD3 + ) and B cell (B220 + ) populations in the indicated peripheral lymphoid organs
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Experimental details: Figure 3 Analysis of T-cell populations in RIP1 t-/- mice. ( a ) Absolute T cell (CD3 + ) and ( b ) B cell (B220 + ) numbers in the spleen and lymph nodes of mutant and control mice. * P
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Experimental details: Fig. 5 EPCAM (FL4) expression in a colon-cancer cell-line-spiked sample on day 3 of storage. Colon cancer cells from HCT-116 cancer cell line were spiked into whole blood from a healthy donor. After sample staining with CD45, PECAM1 and EPCAM antibodies, EPCAM positive cells were selected by excluding CD45 and PECAM1 positive cells during gating on FCSExpress V3
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Supportive validation

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Experimental details: Fig. 6 Dot plots showing a expression of CD227 (FL1) and b expression of EPCAM (FL4) in a breast-cancer cell-line-spiked sample on the day 2 of storage. Breast cancer cells from MDA-MB-231 cancer cell line were spiked into whole blood from a healthy donor. After sample staining with CD45, PECAM1, EPCAM and CD227 antibodies, EPCAM and CD227 positive cells were selected by excluding CD45 and PECAM1 positive cells during gating on FCSExpress V3
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Supportive validation

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Experimental details: Fig. 1 MCL-1 represents amongst the BCL-2 proteins the most promising therapeutic target in AML. a Primary AML samples were treated with indicated concentrations of ABT-199, A1331852 or S63845 for 20 h. Cell death was measured by FACS analysis using Annexin-V/FITC staining in combination with anti-CD45/APC staining. Mean of each experiment performed in duplicate or triplicate is shown. b Eleven AML cell lines were treated with indicated concentrations of ABT-199, A1331852 or S63845 for 72 h prior to analysis of viability using CellTiter-Glo. Data shown are mean and standard error of mean ( n = 3-7).
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Supportive validation

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Experimental details: Fig. 3. Administration of anti-DKK2 antibody enhanced cytotoxicity of NK and CD8 + T cells. (A-F) Flow cytometry analysis of tumor-in-filtrating lymphocytes in Kras G12D/+ ; Apc fl/fl mice. Leukocytes from tumors from Kras G12D/+ ; Apc fl/fl mice injected with of 5F8 or IgG (10 mg/Kg, i.p.) were prepared and analyzed by flow cytometry. (A) percentage of CD45 was shown. (B) CD4, CD8, NK were pre-gated from CD45 + population. (C) Granzyme B expression was measured in both CD8 and NK cells. (D) additional markers of tumor infiltrated NK1.1 + cells were described. (E) Flow cytometry analysis of additional markers of tumor infiltrated CD8 + T cells. Data are presented as means +- sem (Two-sided Student's t-test; n = 5) (*P < 0.05; **P < 0.01; ***P < 0.001).
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Supportive validation

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Experimental details: Figure 1 Identification of dental pulp stem cells (DPSCs). Human DPSCs were positive for the cell surface antigens CD73, CD90, and CD105, as well as negative for CD14, CD20, CD34, and CD45 demonstrated by flow cytometry ( A ). DPSCs were cultured under osteogenic ( B , 14 days) or adipogenic ( C , 21 days) conditions, and showed mineralized nodules and lipid clusters as revealed by alizarin red and oil red staining, respectively. Scale bar = 400 ( B ) or 100 ( C ) mum.
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Experimental details: Figure 2 Identification and purification of hPDLSCs by flow cytometry. hPDLSCs, human periodontal ligament stem cells.
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Experimental details: Figure 1 Identification of ADSCs. ( A ) ADSCs were isolated from the adipose tissue of breast tumors. After two to three passages, the expressions of ADSCs markers (CD90FITC, CD105PE, and CD44FITC) and the lack of CD34PE and CD45FITC were confirmed by flow cytometry. ( B ) The differentiation ability of ADSCs was tested by adipogenesis, osteogenesis, and chondrogenesis.
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Experimental details: Fig. 4 Silence of Usp47 inhibits BCR-ABL T315I -induced CML and inhibits proliferation of KBM T315I in a xenograft model. a Survival of mice after the transplantation of BCR-ABL T315I -transduced Usp47 +/+ or Usp47 -/- BM cells ( n = 6 biologically independent samples per group). **** p < 0.0001, p -value was analyzed by Mantel-Cox-log-rank test. The experiment was repeated three times. b , c Size ( b ) and weight ( c ) of the spleen in the two groups of mice ( n = 5 biologically independent samples per group) at 28 days after transplantation. Center lines of the box and whisker plot represent median values, whereas the box edges indicate the 25th and 75th centiles, and the whiskers indicate the minimum and maximum values. d , e At 28 days after transplantation, the morphology of cells from PB ( d ) was examined by Wright-Giemsa staining; the percentages of GFP + and Gr-1 + cells ( e ) in BM were examined by FACS analysis. f H&E staining of the liver and spleen at 28 days after transplantation. Scale bars, 50 mum. g Ctrl shRNA or ShUSP47-2 transfected KBM5 T315I cells ( n = 8 biologically independent samples per group) were injected into B-NDG mice through the tail vein. **** p < 0.0001, p- value was analyzed by Mantel-Cox-log-rank test. h Size and weight of spleen at day 21 ( n = 6 biologically independent samples per group). Center lines of the box and whisker plot represent median values, whereas the box edges indicate the 25th and 75th centiles, and the whiskers indicate
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Supportive validation

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Experimental details: 10.1371/journal.pone.0235705.g004 Fig 4 Flow cytometry analysis of the COV434 organoids after lymphocyte infiltration assay. COV434 organoids were dissociated and the infiltrated lymphocytes were stained with different markers and analyzed by flow cytometry as described in 'Materials and Methods'. Panel A and B are showing the status of lymphocytes infiltrated in DMSO control organoids and SP-2577 treated organoids, respectively, in the representative dot plots. (Ai and Bi) The dot plots showed the CD3 + population in CD45 + gated lymphocytes. (Aii and Bii) The status of CD4 + and CD8+ T cells gated on CD3 + cells are presented in different quadrants. (Aiii and Biii) Percentage of CD56+ NK in CD3+ cells are presented with quadrant statistics. (Aiv and Biv) The status of CD19+ B cells are shown in CD3+ cells.
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Supportive validation

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Experimental details: Figure 3. SARS-COV-2 infection increases granulocytes and inflammatory monocytes in the bronchoalveolar lavage fluid. K18-hACE2 C57Bl/6J mice were intranasally inoculated with 8 x 10 4 TCID 50 of SARS-CoV-2 Hong Kong/VM20001061/2020 (Source: BEI Resources). Bronchoalveolar lavage was collected, cells isolated, and stained via flow cytometry. * P < 0.05. A two-tailed Student's t test was used to determine statistical significance.
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Supportive validation

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Experimental details: Fig. 3 Hepatic resident-like CD8 + PD1 + T cells are increased in livers of patients with NAFLD patients. a , b , UMAP representation showing the FlowSOM-guided clustering of CD45 + cells ( a ) and flow cytometry plots ( b , left) and quantification ( b , right) of CD8 + PD1 + CD103 + cells derived from hepatic biopsies of healthy individuals or patients with NAFLD or NASH (Supplementary Table 2 ). Populations in b : violet, CD8 + ; red, CD8 + PD1 + CD103 + . T reg cells, regulatory T cells. c , UMAP representations and analyses of differential gene expression by scRNA-seq of CD3 + cells from control individuals or patients with NAFLD or NASH. MAITs, mucosal-associated invariant T cells. d , Correlation of significant differentially expressed genes in liver-derived CD8 + PD1 + T cells compared to CD8 + PD1 - T cells from mice fed with CD-HFD for 12 months and patients with NAFLD/NASH. Shading shows 95% CI. e - h , Expression ( e ) and transcriptional activity ( f ) of velocity analyses of scRNA-seq data, and gene expression ( g ) and correlation ( h ) of expression along the latent-time of selected genes along the latent-time of liver-derived CD8 + T cells from patients with NAFLD or NASH in comparison to control or NASH mouse liver-derived CD8 + T cells. Root cells: yellow, root cells; blue, cells furthest from the root by RNA velocity. End points: yellow, end-point cells; blue, cells furthest from defined end-point cells by RNA velocity. Latent time (pseudo-time by RNA velo
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Experimental details: Figure 6 Gating strategy Flow cytometric gating strategy to sort viable CD45 + CD3 + CD4 + CCR7 - CD45RA - cells from fetal intestinal lamina propria.
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Experimental details: Figure 2 Separation of CTCs and leukocyte or endothelial cells. ( A ) Technical concept using CTC-FIND for single cell isolation. ( B ) Scheme of cell classification. ( C - E ) Representative fluorescence image of CTCs detected in clinical blood specimens. The blood samples (8 mL) collected from patients with mCRC were evaluated by fluorescence microscopy after separation to single cells by CTC-FIND. Specimens were enriched and stained with Hoechst 33342 (pseudo colored blue) and negative markers composed of anti-CD45 and anit-CD50 antibodies (pseudo colored red), followed by the acquisition of fluorescence images for each marker and digital overlay of the two images (merge). Hoechst 33342-positive and CD45/50-negative CTCs were counted and collected from each specimen. Gallery of CTCs ( C ), CTC clusters ( D ) and leucocytes ( E ). Original magnification, x500.
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Experimental details: Fig. 2 Effect of E2 supplementation on immune humanization in mice. a Experimental timeline. 6-7 weeks old NSG-SGM3 female mice were injected with 20 mg/kg/day busulfan on two consecutive days. Five days later, mice were IV injected with 85,000 hCD34 + HSCs and 24 h later received E2 supplemented acidified drinking water for 3.5 weeks ( n = 5, HSC donor ID 7734, KIR-mismatched to HCI-013). Control mice were kept on unsupplemented acidified drinking water. Mice were harvested 13.5 weeks post-humanization, at which point BM and blood were freshly isolated, and flow staining was performed. Flow cytometry was performed on the day of tissue harvest without fixation. b Flow cytometry analysis of hCD45 + cells in BM, and blood of humanized mice receiving E2 or unsupplemented acidified drinking water are shown as percent of viable cells ( n = 2-4). c Flow cytometry analysis of hCD11b + , hCD3 + , hCD56 + and hCD19 + cells in BM and blood of humanized mice receiving E2 or unsupplemented acidified drinking water are shown as percent of hCD45 + cells ( n = 2-4)
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Supportive validation

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Experimental details: Fig. 4 Immune humanization of mice bearing HCI-013 PDX tumors without exogenous E2. a Experimental timeline. 6-7 weeks old female NSG-SGM3 mice were injected with 20 mg/kg/day busulfan on two consecutive days. Five days later, mice were IV injected with 85,000 hCD34 + HSCs and 24 h later implanted with HCI-013 tumor fragments into the MFP. Animals were harvested 18.5 weeks post-humanization ( n = 10, HSC donor ID BM4410, KIR-matched to HCI-013). Control tumors were implanted into non-humanized NSG-SGM3 mice ( n = 3). On the day of harvest, BM, blood and spleens were isolated, and flow staining and flow cytometry were performed on the same day without fixation. b Flow cytometry analysis of hCD45 + cells in BM, blood and spleen of humanized mice are shown as percent of viable cells ( n = 8). c Flow cytometry analysis of hCD3 + , hCD11b + , hCD19 + and hCD56 + cells in BM, blood and spleens of humanized mice are shown as percent of hCD45 + cells ( n = 8). All data are shown as mean +- SEM
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Experimental details: Hemangioma-derived mesenchymal stem cells (Hem-MSCs) express CD103 and CD90, but do not express CD34 or CD45, according to flow cytometry.
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Experimental details: Figure 3 Retinoic Acid Inhibition Increases the Differentiation Capacity of Hematopoietic Progenitors toward Myeloid and Lymphoid Lineage (A) CFU potential of 500 sorted progenitors (CD45/CD43 + CD34 + ) from DEAB-treated cultures (green) and DMSO control (gray) from four independent experiments (n = 4). CFU potential of progenitors similarly sorted from cord blood (CD45/CD43 + CD34 + ) is shown in red. (B) Lymphoid/myeloid differentiation assay performed on iPSC-derived hematopoietic cells. FACS plots and bar graphs showing non-myeloid (CD33/CD14 - ), CD19 - , CD56 + , and CD56 + CD16 + NK fraction from wells seeded with 50 HSC-like cells (CD45/43 + CD34 hi CD38 lo CD90 + CD45RA - ) (17 individual wells) cultured on OP9 stroma for 4 weeks. Left bar graph shows the cluster size of the NK fraction. Right bar graph shows the frequency of this population in the viable non-stroma fraction. See also Figure S2 . (C) FACS plots showing non-myeloid, CD7 + , CD45RA + multipotent early lymphoid progenitors from 3,500 sorted CD34 + cells cultured on OP9DL1 stroma for 3 weeks from one out of two independent experiments (n = 2). FACS plots are gated on the viable non-stroma fraction. Bar graph show the total output of differentiated cells from progenitors generated with DMSO, DEAB, or RA. FACS gates are based on FMO controls. Data represent mean +- SEM. Asterisks indicate significant differences ( * p < 0.05).
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Supportive validation

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Experimental details: Figure 5 Decreased Retinoic Acid Signaling Maintains the Primitiveness of Generated Hematopoietic Progenitors (A) Representative FACS profiles of cells harvested at day 16 and after extended culture period at day 21. Cells were treated with or without DEAB at 10 muM during the whole culture period. (B) Graphs showing fold changes in maturation of the hematopoietic population (CD45/43 + ) observed between day 16 and day 21 for cells treated with DEAB or DMSO from three independent experiments (n = 3). Data represent mean +- SEM. Asterisks indicate significant differences ( * p < 0.05).
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Supportive validation

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Experimental details: Figure 1 Primary human adipose-derived cells cultured in hypoxia (hAMSCs-H) and normoxia (hAMSCs-N) are both MSCs but normoxia-cultured cells show increased signs of senescence, such as increased area and elongated morphology, compared with hypoxia-cultured cells. ( a ) hAMSCs were isolated from human fat tissue and cultured in hypoxic (1.5% oxygen) or normoxic (21% oxygen) conditions in vitro . The viability, mobility, tumor tropism, safety, and tumorigenic potential were subsequently compared in vitro and in vivo . ( b ) Differentiation assay. hAMSCs were cultured in control media and differentiation media for 3 weeks, 10 days after the second passage. Three different stains were performed to assess differentiation capabilities (scale bar, 100 mu m). ( c ) Flow cytometric analysis was performed to confirm the absence of CD31-, CD34-, and CD45-positive cells in both cell cultures. In addition, primary hAMSC cultures expressed high levels of CD73, CD90, and CD105, both in hypoxic and normoxic culture conditions at day 10 after passage 2. ( d ) Representative images of cell morphologies of hAMSCs on 2D surface (scale bar, 200 mu m). ( e ) Schematic of 3D-nanopatterned surface used to assess morphology and motility. ( f ) Images of cell morphologies of hAMSCs on 3D-nanopatterned surface (scale bar, 200 mu m). ( g - j ) The length, width, area, and length-to-width ratio were measured and compared after cell aligned on the nanopattern surface. Error bars represent S.E.M. * P
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Supportive validation

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Experimental details: Figure 3 Hypoxia-cultured primary human adipose-derived mesenchymal stem cells (hAMSCs-H) retain a greater proliferation capacity compared with normoxia-cultured primary hAMSCs (hAMSCs-N) when exposed to GBM media. hAMSCs-H maintain stem cell characteristics when exposed to GBM media. ( a ) Representative MRI of GBM from a patient. ( b ) Schema showing the collection of GBM CM and culture of hAMSCs in filtered GBM CM for proliferation and migration assays. ( c ) MTT assay was used to determine the effects of hypoxic conditions on the proliferative capacity of primary hAMSCs in GBM CM. In GBM CM, hAMSCs-H showed greater proliferation at day 10 and 15 compared with hAMSCs-N. ( d ) Ki-67 immunostaining was performed to quantify the number of proliferating cells in GBM CM. Proliferative capacities of hAMSCs-H and hAMSCs-N are shown in GBM CM (normalized to hAMSC-N proliferative capacity in control media). In GBM CM, hAMSCs-H had a greater proportion of proliferating cells than hAMSCs-N. ( e) Differentiation assay. hAMSCs were cultured in control media, differentiation media, and GBM CM for 3 weeks, 10 days after the second passage. Three stainings were performed to assess the differentiation capabilities (scale bar, 100 mu m). Both hAMSCs-N and hAMSCs-H maintained tri-lineage differentiation capability in GBM CM. ( f ) Flow cytometric analysis for CD31, CD34, CD45, CD73, CD90, and CD105 in hAMSC-N and hAMSC-H cultures after exposure to GBM CM for 20 days. hAMSCs-H maintained MSC
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Supportive validation

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Experimental details: Figure 1 Detection of cells with abnormal nuclei using imaging flow cytometry. ( a ) Representative cell images with the same DAPI intensity. CD45, a lymphocyte biomarker, was labeled with PE-Cy5. Upper panel: CD45 + lymphocyte with a smaller DAPI area. Lower panel: CD45 - cells with a larger DAPI area, showing a looser structure of nuclei. Scale bars represent 20 mum. ( b ) Mean DAPI fluorescence intensity in PBMCs. Unlike the cells with a normal DAPI area (lower panel), a G1 and G2 peak could be detected in the cells with larger DAPI areas (upper panel). ( c ) Imaging flow cytometry test results for the peripheral blood samples from MVI patients. Left panel, basic results for peripheral blood mononuclear cells (PBMCs), the horizontal axis indicates the cell area and the vertical axis indicates the aspect ratio. The gate displays the group of single cells. Right panel: patients with MVI had 68.1 +- 14.8 cells with large DAPI areas. ( d ) Representative images for PBMCs from HCC patients. CD45, a lymphocyte biomarker, was labeled with PE-Cy5. EpCAM, a biomarker of circulating tumor cells (CTCs), was labeled with FITC. Left panel: cells with high karyoplasmic ratios (HKR cells), which were EpCAM-positive and CD45-negative. Right panel: cells with normal karyoplasmic ratios (normal cells), which were EpCAM-negative and CD45-positive. Scale bars represent 20 mum. ( e ) Flow cytometry test results for HKR cells from MVI patients. All HKR cells were CD45 - , but only 8.7% of HKR c
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Supportive validation

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Experimental details: Figure 3 Diagnostic value of HKR cell content. ( a ) Youden curve for HKR cells in 52 HCC patients and 12 normal individuals. Arrow refers to the maximum Youden index, which was defined as the threshold. ( b ) ROC curve of HKR cells for 23 HCC patients with MVI (M1 & M2) and 18 HCC patients without MVI (M0). HKR cells were identified by imaging flow cytometry. ( c ) Relative number of CTCs in 100,000 peripheral blood mononuclear cells in non-cancer liver disease patients and HCC patients. CTCs were defined as CD45 - & EpCAM + cells in the peripheral blood and detected by imaging flow cytometry. Data are presented as means +- SD of independent experiments. ( d ) Relative number of CTCs in 100,000 peripheral blood mononuclear cells in HCC patients with MVI (M1 & M2) and HCC patients without MVI (M0). CTCs were defined as CD45 - & EpCAM + cells in peripheral blood and detected by imaging flow cytometry. Data are presented as means +- SD of independent experiments. * P < 0.05. ( e ) ROC curve of CTCs for 23 HCC patients with MVI (M1 & M2) and 18 HCC patients without MVI (M0) in 52 total samples. CTCs were defined as CD45 - & EpCAM + cells in peripheral blood and detected by imaging flow cytometry.
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Supportive validation

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Experimental details: Figure 5 Inefficient Hematopoietic Outcomes from HEB -/- hESC-Derived Progenitors (A) Experimental overview for the colony-forming unit (CFU) assay of erythromyeloid potential. Cell suspensions from day-18 (d18) EBs were obtained and placed into methylcellulose cultures either as an unfractionated population (top) or as four sorted subsets based on combinatorial expression of CD34 and CD45 (bottom). (B) Numbers of erythroid (BFU-E) and myeloid (CFU-GM) arising from unfractionated WT and HEB -/- d18 EB-derived cells in methylcellulose cultures. (C) Flow-cytometric analysis for CD34 and CD45 expression of unfractionated WT and HEB -/- d18 EB-derived cells. (D and E) The percentages (D) and cell numbers (E) of CD34 + CD45 - , CD34 + CD45 + , and CD34 - CD45 + subsets within WT and HEB -/- d18 EBs. (F) Numbers of erythroid (BFU-E) and myeloid (CFU-GM) arising from CD34 + CD45 + cells sorted from WT and HEB -/- d18 EBs. (G) Experimental overview for T cell differentiation. CD34 + cells were sorted from d8 EBs and co-cultured with OP9-DL4 cells. (H and I) Percentages (H) and numbers (I) of CD45 + cells at successive days (d) of OP9-DL4 co-cultures. Error bars represent mean +- SD (n = 3 independent experiments). *** p < 0.005 by Student's t test. Plots in (C) are representative of three independent experiments. See also Figures S5 and S6 .
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Supportive validation

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Experimental details: Figure 6 Ectopic Expression of HEBCan in HEB -/- hESCs Restores Lineage-Specific Gene Expression and Hematopoietic Specification (A) Western blot analysis for HEB expression in WT, KO (HEB -/- ), KO + GFP (HEB -/- hESCs transduced with GFP control vector) and KO + HEBCan (HEB -/- hESCs transduced with HEBCan-encoding vector) hESCs. (B) Bright-field (top) and fluorescent (bottom) images of day-8 (d8) EBs derived from HEB -/- hESCs transduced with control or HEBCan-expressing lentiviral particles. Scale bar, 100 mum. (C and D) qRT-PCR analysis for the expression of pluripotency-associated genes (C) and mesoendodermal genes (D) in WT, KO + GFP, and KO + HEBCan hESC-derived cells at d0 and d4 of EB culture. mRNA levels are shown relative to GAPDH. (E) Flow-cytometric analysis of CD34 and KDR, CD144, and CD31 on WT, KO + GFP, and KO + HEBCan d8 EB-derived cells. (F and G) Percentages (F) and numbers (G) of CD34 + cells in WT, KO + GFP, and KO + HEBCan d8 EBs. (H) Flow-cytometric analysis for CD34 and CD45 on WT, KO + GFP, and KO + HEBCan d18 EB-derived cells. (I and J) Percentages (I) and numbers (J) of CD34/CD45 subsets in WT, KO + GFP, and KO + HEBCan d18 EB-derived cells. (K) Numbers of erythroid (BFU-E) and myeloid (CFU-GM) arising from unfractionated WT, KO + GFP, and KO + HEBCan d18 EBs. Error bars represent mean +- SD (n = 3 independent experiments). * p < 0.05, ** p < 0.01, *** p < 0.005 by Student's t test. Images in (A) and (B) and plots in (E) and (H) are represent
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Experimental details: Figure 1--figure supplement 1. Human cell reconstitution and HIV-1 infection in BLT hu-mice. ( A ) Representative flow analysis plots showing side scatter (SSC-A) and forward scatter (FSC-A), human CD45+ hematopoietic cell population, human CD3+ T lymphocyte and CD19+ B lymphocyte, and human CD4+ T and CD8+ T lymphocyte populations in peripheral blood at 13 weeks post-BLT transplant. ( B ) Percentage of reconstituted human hematopoietic cell populations in peripheral blood in all BLT mice. ( C ) Schematic for infection and analysis of BLT hu-mice. Animals were euthanized at days 0, 5, 10, 63, and 92 post-HIV-1 injection for analysis of blood and bone marrow. ( D ) Human CD4/CD8 ratios were monitored in peripheral blood by flow cytometry. ( E ) Quantification of HIV-1 viral load in peripheral blood at the time of euthanasia.
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Supportive validation

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Experimental details: Figure 6 NK cells in the livers of NAFLD patients are less cytotoxic than those from healthy controls. (A) Immune cells were isolated from human livers. NK cells were identified by scatter, and as live CD45 + CD56 + CD3 - Tbet hi Eomes lo cells. Liver-resident NK cells (lrNK) were identified as live CD45 + CD56 + CD3 - Tbet lo Eomes hi cells. (B,C) Intrahepatic leukocytes were cultured for 4 h in the presence of anti-CD107a and Brefeldin A. Representative CD107a staining of NK cells from a healthy (B) and a NAFLD (C) liver is shown. (D) Inverse correlation between NK cell degranulation and histological score in human livers. H, healthy; SS, simple steatosis; NAS, NASH Activity Score; C, cirrhosis. Significance was determined using Spearman's Rank Correlation. (E) CD56 + CD3 - CD16 + CXCR6 - NK cells were sorted from healthy ( n = 4) and NAFLD ( n = 3) livers and cultured for 4 h with the human NK target cell, K562. At the end of culture, target cell death was assessed using AnnexinV staining. (F) Inverse correlation between liver-resident NK cell degranulation and histological score in human livers. Significance was determined using Spearman's Rank Correlation. (G) CD56 + CD3 - CXCR6 + liver-resident NK cells were sorted from healthy ( n = 4) and NAFLD ( n = 3) livers and cultured for 4 h with the human NK target cell, K562. At the end of culture, target cell death was assessed using AnnexinV staining. For (E,G) , medians are shown. For these small datasets, no statistical te
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Supportive validation

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Experimental details: Figure 1. Characterization of hUC-MSCs. (A) Morphological observations of hUC-MSCs. Umbilical cord tissues were cultured for >15 days and long spindle-shaded fibroblastic cells were observed around the tissue using Zeiss light microscopy (scale bar, 100 um). (B) Phenotyping of hUC-MSCs. hUC-MSCs were stained with a fluorescein-labeled antibodies (CD34, CD45, CD73, CD90, CD105, CD14, CD19 and HLA-DR) and analyzed with a flow cytometer. (C) Adipogenic and (D) osteogenic differentiation of hUC-MSCs. hUC-MSCs were cultured in adipogenic and osteogenic medium, respectively. Lipid droplets in the adipocytes are presented with Oil Red O staining (scale bar, 100 um). hUC-MSCs-derived osteoblasts were detected with Alizarin Red staining (scale bar, 200 um). (E) hUC-MSCs inhibit the proliferation of CFSE-labeled CD4 + T cells, which were activated by Con A stimulation. Experiments were repeated three times and representative graphs and images are presented. hUC-MSC, human umbilical cord-derived mesenchymal stem cell; MSC Sup, culture supernatant of hUC-MSCs; Con A, concanavalin A; CFSE, carboxyfluorescein succinimidyl ester.
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Supportive validation

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Experimental details: Figure 4 Increased accumulation of fibrocytes and CCR5-positive cells in the dermis of SSc patients . Fibrotic and control human skin sections were stained using the indicated antibodies and DAPI to detect nuclei. Large numbers of double-positive cells were observed in SSc dermis for each comparison ( A , CCR5 vs. Procol I; C , Procol I vs. CD45; E , CCR5 vs. CD45), while almost none were observed in Control dermis. These representative fields were photographed at 400 x magnification. To quantify these observations, the number of double-positive cells was counted in three subjects per category, five high power fields (HPF) per subject. The data are presented in terms of the number of double-positive cells per HPF (average +- s.e.m.). (B) quantification of (A) ; (D) quantification of (C) ; (F) quantification of (E) .
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Supportive validation

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Experimental details: Figure 5 Increased accumulation of fibrocytes and CCR5-positive cells in fibrotic mouse dermis is reversed by CSD . Mice were treated with bleomycin and CSD as described in Figure 3 . Skin sections were stained using the indicated antibodies and DAPI to detect nuclei. Large numbers of double-positive cells were observed in bleomycin-treated mice (that did not receive CSD) for each comparison ( A , CCR5 vs. Procol I; B , Procol I vs. CD45; C , CCR5 vs. CD45), while almost none were observed in Control mice. CSD treatment almost completely blocked the accumulation of these cells. These representative fields were photographed at 400 x magnification. To quantify these observations, the number of double-positive cells was counted in three subjects per category, five HPF per subject. The data are presented in terms of the number of double-positive cells per HPF (average +- s.e.m.). (D) quantification of (A) ; (E) quantification of (B) ; (F) quantification of (C) .
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Supportive validation

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Experimental details: Figure 6 Increased accumulation of CCR5 ligand-positive cells in the dermis of SSc patients . Fibrotic and control human skin sections were stained using the indicated antibodies and DAPI to detect nuclei. Large numbers of double-positive cells were observed in SSc dermis for each comparison ( A , MIP1alpha vs. CD45; C , MIP1beta vs. CD45), while almost none were observed in Control dermis. These representative fields were photographed at 400 x magnification. To quantify these observations, the number of double-positive cells was counted in three subjects per category, five HPF per subject. The data are presented in terms of the number of double-positive cells per HPF (average +- s.e.m.). (B) quantification of (A) ; (D) quantification of (C) .
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Supportive validation

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Experimental details: Figure 7 Increased accumulation of CCR5 ligand-positive cells in fibrotic mouse dermis is reversed by CSD . Mice were treated with bleomycin and CSD as described in Figure 3 . Skin sections were stained using the indicated antibodies and DAPI to detect nuclei. Large numbers of double-positive cells were observed in bleomycin-treated mice (that did not receive CSD) for each comparison ( A , MIP1alpha vs. CD45; B , MIP1beta vs. CD45), while almost none were observed in Control mice. CSD treatment almost completely blocked the accumulation of these cells. These representative fields were photographed at 400 x magnification. To quantify these observations, the number of double-positive cells was counted in three subjects per category, five HPF per subject. The data are presented in terms of the number of double-positive cells per HPF (average +- s.e.m.). (C) quantification of (A) ; (D) quantification of (B) .
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Supportive validation

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Experimental details: 1 Figure Representative photomicrographs of BMSCs on Day 7 of tertiary culture in (a) negative immune reaction of anti-CD45 antibody in a negative control dish. (b) Showing most of the cells exhibit positive brownish immune reaction for anti-CD44 antibody (arrows). Phase contrast photomicrographs: x200. BMSC, bone marrow mesenchymal stem cell
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Supportive validation

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Experimental details: Figure 1 T-cell infiltration in normal and tumor tissues in primary breast cancer ( A ). Representative images of immunohistochemical staining of tumor-infiltrating CD3 + T cells in formalin-fixed paraffin embedded breast non-tumor (NT) and tumor tissues (TT). ( B ). Freshly isolated immune cells infiltrating NT (NILs) and TT (TILs) from 11 PBC patients were stained with 7AAD, CD45, CD3, CD4 and CD8 antibodies for identification of T cells and their subsets. Representative flow cytometric plots of surface staining from one cancer patient are shown. 7AAD dye was used to gate live cells, followed by lymphocyte identification by CD45 and CD3 stainings. Different subsets of T cells were then characterized using CD4 and CD8 antibodies. ( C ). Scatter plots showing the differences in tissue-infiltrating CD45 + CD3 - , CD45 + CD3 + , CD4 + and CD8 + cells between NILs and TILs.
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