Characterization and Clinical Application of Human CD34+ Stem/Progenitor Cell Populations Mobilized into the Blood by Granulocyte Colony-Stimulating Factor



A phase I study was performed to determine the safety and tolerability of injecting autologous CD34+ cells into five patients with liver insufficiency. The study was based on the hypothesis that the CD34+ cell population in granulocyte colony-stimulating factor (G-CSF)-mobilized blood contains a subpopulation of cells with the potential for regenerating damaged tissue. We separated a candidate CD34+ stem cell population from the majority of the CD34+ cells (99%) by adherence to tissue culture plastic. The adherent and nonadherent CD34+ cells were distinct in morphology, immunophenotype, and gene expression profile. Reverse transcription-polymerase chain reaction-based gene expression analysis indicated that the adherent CD34+ cells had the potential to express determinants consistent with liver, pancreas, heart, muscle, and nerve cell differentiation as well as hematopoiesis. Overall, the characteristics of the adherent CD34+ cells identify them as a separate putative stem/progenitor cell population. In culture, they produced a population of cells exhibiting diverse morphologies and expressing genes corresponding to multiple tissue types. Encouraged by this evidence that the CD34+ cell population contains cells with the potential to form hepatocyte-like cells, we gave G-CSF to five patients with liver insufficiency to mobilize their stem cells for collection by leukapheresis. Between 1 × 106 and 2 × 108 CD34+ cells were injected into the portal vein (three patients) or hepatic artery (two patients). No complications or specific side effects related to the procedure were observed. Three of the five patients showed improvement in serum bilirubin and four of five in serum albumin. These observations warrant further clinical trials.


Liver transplantation is the only current therapeutic modality for liver failure, but due to the shortage of organ donors, it is available to only a small proportion of patients. Adult stem cell therapy could solve the problem of degenerative disorders, including liver disease, in which organ transplantation is inappropriate or there is a shortage of organ donors. This view is predicated upon the evidence that stem cells, particularly those in hematopoietic tissue, have the ability to develop into endodermal, mesodermal, and ectodermal cell types [1].

Several sources of stem cells have been proposed as sources for cell therapy. Embryonic stem cells are the most potent in terms of their differentiation potential but may be tumorigenic when transplanted in vivo, and their use is beset by ethical issues [2, 3]. Adult stem cells may be found in any tissue [1], but hematopoietic tissue is most accessible. Hematopoietic tissue contains two types of stem cells, the mesenchymal and hematopoietic stem cells. Mesenchymal stem cells (MSCs) were first described by Friedenstein et al. in 1974 [4]. More recently, MSC cultures were shown to contain multipotential adult progenitor cells (MAPCs) [5], unrestricted somatic stem cells (USSCs) [6], and rapidly self-renewing (RS) cells [7, 8], all of which may represent early stages in MSC development. However, none of these cell types can be identified in fresh hematopoietic tissue [9]. In addition, they cannot be discriminated phenotypically from the bulk MSC population in culture, although Smith et al. [10] have separated RS cells from more mature cells in MSC cultures by flow cytometry based on the forward and side light scattering properties of the cells.

Stem cells in hematopoietic tissue have been used for hematological reconstitution for many years [11]. These cells are CD34+ and CD133+ and give rise to all lineages of blood cell differentiation. Thus, they have the advantage that they can be prospectively isolated from hematopoietic tissue in known numbers. Recently, they have been used to transplant patients with liver disease [12] or cardiac insufficiency [13]. These experiences led us to hypothesize that the regenerative activity of CD34/CD133+ stem cells resided in a CD34+ subpopulation. Here, we sought to identify and characterize the CD34+ cells with regenerative potential and found that they could be separated from the majority of the CD34+ cells by their plastic adherent properties. As we will show, these cells have a small lymphocyte-like morphology that has long been associated with primitive stem cell populations. In light of the knowledge that the CD34+ cell population contained an identifiable candidate stem cell population, we used CD34+ cells for a phase I study of cell therapy in patients with liver disease.

Materials and Methods

Preclinical In Vitro Studies

Cell Source.

Granulocyte colony-stimulating factor (G-CSF)-mobilized peripheral blood cells were obtained from leukaphereses processed by the Stem Cell Laboratory, Hammersmith Hospital, in excess of clinical requirements. Informed consent and local research ethics committee approval were granted in all cases.

Cell Isolation.

CD34+ cells were diluted at 1:4 in Hanks' buffered saline solution (HBSS; Gibco, Paisley, U.K., before the mononuclear cells (MNCs) were separated by centrifugation over a Lymphoprep (Axis-Shield, Kimbolton, Cambridgeshire, U.K., density gradient at 1,800 rpm for 30 minutes (Heraeus, Hanau, Germany, The MNC fraction was collected and washed first in HBSS, then with MACS (magnetic cell sorting) buffer (phosphate-buffered saline buffer supplemented with 0.5% bovine serum albumin and 5 mM EDTA, pH 7.2). CD34+ cells were isolated from MNCs, using the CD34+ positive cell selection kit (MiniMacs; Miltenyi Biotec, Bergisch Gladbach, Germany,

Cell Culture.

Isolated CD34+ cells were plated on 35-mm2 Petri dishes in α-minimal essential medium (α-MEM) supplemented with 15% fetal bovine serum (FBS) and incubated for 2 hours at 37°C and 5% CO2. After 2 hours, the nonadherent cell fraction was removed by washing the plates three times. Adherent CD34+ cells were cultured in α-MEM supplemented with 30% FBS and cytokines (20 ng/ml stem cell factor [SCF], 1 ng/ml GM-SCF, 5 ng/ml IL-3, and 100 ng/ml G-CSF) at 37°C in 5% CO2 in air.

Telomerase Assay.

Cells were lysed in 1× 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid (CHAPS) buffer, and the lysates were analyzed using the TRAPeze telomerase detection kit (InterGen, Burglington, MA, according to the manufacturer's instructions.

RNA Isolation.

RNA was isolated from fresh cells and from cells that had been cultured for 7 days using the RNeasy Mini kit according to the manufacturer's instructions (Qiagen, Crawley, West Sussex, U.K., To ensure purity of the RNA, the samples were treated with DNase (Promega UK, Southampton, U.K., The RNA was then purified using the Qiagen PCR Purification Kit according to the manufacturer's instructions. Total RNA concentration was determined by measuring the optical density at 260 nm in a spectrophotometer (Eppendorf UK Limited, Cambridge, U.K.,

Reverse Transcription-Polymerase Chain Reaction

Reverse transcription-polymerase chain reaction (RT-PCR) was carried out using the One-step RT-PCR Kit (Qiagen). A master mix sufficient for 40 reactions was prepared in a 1.5-ml Eppendorf tube consisting of 300 μl of 5× RT-PCR buffer, 60 μl of dNTP, 150 μl of Q solution, 650 μl of RNase-free water, and 80 μl of RT-enzyme mix. Twenty microliters from the RT-PCR master mix was then aliquoted into each tube, with an overlay of 50 μl of mineral oil to prevent evaporation.

Positive controls were a pool of RNA known to be positive for the genes of interest by RT-PCR and sequence verification by the Medical Research Council sequencing laboratory at the Hammersmith Hospital. Both negative controls and culture medium controls were used to exclude false-positive results. As an internal RNA standard, the housekeeping gene GAPDH (glyceraldehyde-3-phosphate dehydrogenase) was used.

RT-PCR conditions used were as follows: RT at 50°C for 1 hour; PCR activation at 95°C for 15 minutes; three-step cycling at 95°C for 1 minute, N°C for 1 minute, and 72°C for 1 minute for 35 cycles; and final extension at 72°C for 10 minutes, where N is the gene-specific RT-PCR annealing temperature.

Nested PCR.

Nested PCR was performed by adding 2 μl of the corresponding RT-PCR product to each PCR tube along with 2 μl of the appropriate nested forward and backward primer mix and 20 μl of nested PCR mix. For the genes α-1 antitrypsin, c-met, vimentin, α-fetoprotein, β-cellulin, and CK19, a separate PCR mix containing 2.5 μl of 50 mM MgCl2 was prepared. Nested PCR conditions used were as follows: 50°C for 1 minute; 95°C for 15 minutes; and 35 cycles of 95°C for 1 minute, N°C for 1 minute, and 72°C for 1 minute, and 72°C for 10 minutes, where N is the gene-specific nested PCR condition temperature.

Gel Electrophoresis.

RT- and nested PCR products were analyzed by agarose gel electrophoresis and visualized under UV light on a transilluminator (Ultra-Violet Products Ltd., Cambridge, U.K.,, and images were captured using a digital camera (Ultra-Violet Products Ltd.) connected to a computer.

Flow Cytometry.

For surface staining, at least 1 × 104 cells were labeled with directly conjugated or unconjugated antibody. A fluorescein isothiocyanate (FITC)-conjugated secondary antibody was added to cells labeled with unconjugated primary antibody. A commercial cell fixation and permeabilization kit (Caltag Laboratories, Burlingame, CA, was used for intracellular staining according to the manufacturer's instruction. The antibodies used were CD34-phycoerythrin (PE) (BD Biosciences, San Diego,, CD54-PE (intercellular adhesion molecule 1 [ICAM-1]; BD Biosciences), CD184 CXCR4 (BD Biosciences), epithelial cell adhesion molecule (EpCAM)-PE (Miltenyi Biotec), anti-mouse-FITC (Dako UK Ltd., Ely, Cambridgeshire, U.K.,, nestin (AbCam, Cambridge, U.K.,, vascular endothelial growth factor receptor 2 (VEGFR2), and vimentin (Dako UK Ltd.). Appropriate isotype controls were included in all cases. Stained cells were analyzed on a FACS Calibur flow cytometer (BD Biosciences).

Clinical Study

Patient Selection.

Patients were recruited with ethics committee approval and according to criteria determined by the Multidisciplinary Treatment (MDT) committee of the Hammersmith Hospital. Informed consent was granted in all cases. Inclusion criteria were the following: age 20–65 years, chronic liver failure, abnormal serum albumin and/or bilirubin and/or prothrombin time, unsuitable for liver transplantation, World Health Organization performance status less than 2, women of child-bearing potential using reliable and appropriate contraception, life expectancy of at least 3 months, and ability to give informed consent. Exclusion criteria were the following: patients aged less than 20 or more than 65 years; liver tumors or history of other cancer; pregnancy or lactation; recurrent gastrointestinal bleeding or spontaneous bacterial peritonitis; active infection, including HIV; and inability to give informed consent.

Prospective patients were admitted for liver function tests, full blood count, coagulation profile, α-fetoprotein levels, computed tomography (CT) scan, visceral angiography, and a Duplex Doppler scan. They were then discharged home pending a discussion by the MDT committee as to their suitability for inclusion in the study.

Included patients were admitted and given subcutaneously 520 μg of G-CSF (Chugai Pharmaceuticals Co., Ltd., Tokyo, daily for 5 days to increase the number of circulating CD34+ cells. Leukapheresis was performed on day 5. The leukapheresis product was transferred to the laboratory, where CD34+ cells were immunoselected using the CliniMacs device (Miltenyi Biotech). The CD34+ cells were then returned to the patient via the hepatic artery or portal vein in the Imaging Department. Patients were discharged after overnight bed rest.

Patients returned to the outpatient clinic on days 7, 15, 30, 45, and 60 after infusion for liver function tests, full blood count, coagulation profile, and α-fetoprotein assay. In addition, on day 60, ultrasound and CT scans of the liver were performed.


Adherent CD34+ Cells Are Distinct from Nonadherent CD34+ Cells

Separation of plastic adherent cells from MNC fractions is a procedure used to initiate MSC cultures. However, the MSCs rapidly outgrow any other cell types that may be present. Here, we purified CD34+ cells and then subfractionated the population into adherent and nonadherent cells. By direct observation and cell counting, the adherent subpopulation comprises approximately 1% of the total CD34+ cell population. Morphologically, the adherent cells were small and lymphocyte-like with a high nuclear/cytoplasmic ratio, whereas the nonadherent cells exhibit a larger blast cell-like morphology (Fig. 1A). Phenotypically, the adherent CD34+ cells were 99% pure and expressed uniformly low levels of CD38, CD33, and HLA-DR. In this characteristic, they differ from the nonadherent CD34+ cells, which exhibit significantly greater variability in the expression of these antigens (Fig. 1B).

Figure Figure 1..

Comparison of adherent and nonadherent CD34+ cells at isolation. (A): Morphologies of freshly isolated adherent (left panel) and nonadherent (right panel) CD34+ cells. (B): Percentages of CD38, CD33, and HLA-DR cells in the adherent and nonadherent CD34+ cell fractions. (C): Polymerase chain reaction analysis of gene expression by adherent and nonadherent CD34+ cells. (D): Flow cytometric analysis of rhodamine staining of adherent and nonadherent CD34+ cells. x-axis, forward scatter; y-axis, fluorescence intensity. Cell within the gate stain brightly with rhodamine.

We used RT-PCR to compare the gene expression profiles of the adherent and nonadherent CD34+ fractions. Fifteen genes were analyzed, and all were expressed in the adherent CD34+ cells, whereas only two were expressed in the nonadherent CD34+ cell fraction (Fig. 1C). These two were angiopoietin-2 and ICAM-2, both of which are associated with hematopoietic cell differentiation [14, 15].

Overall, these data show that the adherent and nonadherent CD34+ cells are distinct populations that can be distinguished by their different morphologies, phenotypes, and gene expression profiles.

Adherent CD34+ Cells Are Primed to Generate Multiple Tissue Types

Enver and colleagues [16] hypothesized that primitive stem cells express genes that indicate their potential to differentiate into different lineages and cell types. Accordingly, we examined the gene expression profile of the cells by RT-PCR using probes corresponding to genes known to be expressed in liver and in a range of other nonhematopoietic tissues. Figure 2A demonstrates the expression of genes associated with liver cell differentiation. As may be seen from Figure 2B, the cells also expressed multiple genes associated with the stem cell state and genes associated with pancreatic, cardiovascular, muscle, and nerve cell differentiation. Expression at the protein level was demonstrated for CD34, VEGFR2, CD54, nestin, and EpCAM by flow cytometry (Fig. 2C–2H).

Figure Figure 2..

Expression analysis of adherent and nonadherent CD34+ cells at isolation. (A): Expression of genes associated with liver cell differentiation by freshly isolated adherent CD34+ cells. Lane 1: Fresh adherent CD34+ cells. Lane 2: Negative control. Lane 3: Culture medium control. (B): Expression of genes associated with stem cells and with pancreatic, cardiovascular, and nerve cell differentiation by freshly isolated adherent CD34+ cells. (C–H): Flow cytometric analysis of freshly isolated CD34+ cells. x-axis, forward scatter; y-axis, fluorescence intensity. Antigen-positive cells are located within the gate. Abbreviation: bp, base pairs.

Although none of the genes tested is exclusive to the stem cell state or tissue lineage differentiation, these results indicate that the adherent CD34+ cell population is a putative stem/progenitor cell population that may have the potential to generate cells corresponding to a range of tissue types.

Diverse In Vitro Differentiation Potential of Adherent CD34+ Cells

Adherent CD34+ cells have been isolated from 100% of the 125 leukapheresis samples analyzed and have been cultured successfully in 98.4% of cases. At the initiation of the culture, the cells did not express telomerase, as assessed by the telomere repeat amplification protocol (TRAP) assay, which is consistent with stem cell quiescence, but telomerase was detectable in 7-day-old cultures (Fig. 3A). During culture, the adherent cells increased in number and released nonadherent cells into the culture supernatant to achieve a 3-log increase in total cell number in 2–3 weeks (Fig. 3B). The morphology of the majority of the adherent cells remained round and mononuclear for the first 3–5 days of culture, and clusters of adherent CD34+ cells were also observed, indicating that the primitive cells can self-renew and maintain their phenotype (Fig. 3C). Morphological differentiation began after day 3, and the proportion of differentiated cells increased with time. The differentiated cells assumed various morphologies, including adherent spindle-shaped fibrobast-like cells (Fig. 3C). In addition, large “colonies” of cells were seen floating in the culture supernatant. These colonies were picked out of the cultures, cytospun onto glass microscope slides, and stained with May Grunwald-Giemsa. They consisted of granulocytic, monocyte-macrophage, megakaryocytic, and erythroid cells (Fig. 3D). The cells were also plated into a standard hematopoietic colony assay. Typically, approximately 1% of the cells formed granulocyte-macrophage colonies (CFU-GM). Burst-forming units-erythroid (BFU-E), megakaryocytic CFU-Mk, and multipotential colony-forming unit-granulocyte, -erythrocyte, -monocyte, and -megakaryocyte (CFU-GEMM) were also seen.

Figure Figure 3..

Characterization of cultured adherent CD34+ cells. (A): Telomerase activity in freshly isolated CD34+ cells and in their progeny in culture. Telomerase activity was not found in fresh cells but was detected in cultured cells. (B): Growth curve of adherent CD34+ cells and their progeny in culture showing cumulative and actual cell numbers. (C): Morphologies of adherent CD34+ cells and their progeny in culture. Top left and center panels show cells immediately after plating and after 3 days of culture, respectively. (D): Morphology of cells recovered from “colonies” floating in the culture supernatant ([C], middle left panel).

Extensive Gene Expression Is Maintained by Cultured CD34+ Cells

The expression of genes associated with liver cell differentiation was maintained in culture (Fig. 4A) as was expression of genes for cardiovascular, muscle, and nerve cell differentiation (Fig. 4B). Expression at the protein level was demonstrated by flow cytometry and immunocytochemistry (Fig. 4C–4K). This maintenance of gene expression by cultured cells indicated that the adherent CD34+ cells may generate progeny capable of differentiating along multiple tissue lineages.

Figure Figure 4..

Expression analysis of cultured adherent CD34+ cells. (A): Expression of genes associated with liver cell differentiation by cultured adherent CD34+ cells. Lane 1: Cultured cells. Lane 2: Negative control. Lane 3: Culture medium control. (B): Expression of genes associated with stem cells and with pancreatic, cardiovascular, and nerve cell differentiation by cultured adherent CD34+ cells. (C–K): Flow cytometry and immunocytochemistry. Abbreviation: bp, base pairs.

Clinical Study

Five patients (four males and one female) aged 49–61 (mean 49) years were recruited to the study and underwent the entire procedure. In three patients the cells were injected into the portal vein under CT scan, and in two patients they were injected via the hepatic artery. The etiology, diagnosis, clinical signs, results of investigations, cell concentrations, and postinfusion observations for each patient are shown in Tables 1, Table 2. to 3.

Table Table 1.. Etiology of patients in phase I trial
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Table Table 2.. Clinical signs and investigations of patients in phase I study
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Table Table 3.. Injection routes and cell concentrations plus postinjection observations
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After G-CSF treatment, increased total white blood cell counts indicated mobilization of progenitor cells in all patients. All patients experienced thrombocytopenia (as expected after leukapheresis), but platelet counts returned to baseline within 1 week. Patients were given 1 × 106–2 × 108 CD34+ cells as a single bolus. No mortalities or specific side effects, except for mild pain and discomfort at the site of CD34+ cell infusion, were recorded. In particular, there was no bleeding or infection of significant deterioration in liver function. CT scans showed no evidence of focal liver lesions in any patient either before or after treatment, and Duplex Doppler ultrasound scans showed patent portal veins with hepatopedal flow.

Patient 1 showed initial improvement in serum bilirubin from 32 mM to normal, but this reverted to baseline by day 60; in patient 2, C-reactive protein (25–7 mg/l) and serum bilirubin (31–15 mM) normalized and serum albumin increased from 33–37 g/l; the only changes recorded in patient 3 were decreased ALT (alanine aminotranferease) from 87–61 U/l and AST (aspartate aminotransferase) from 131–92 U/l); patient 4 experienced an initial improvement in liver function, but this was interrupted due to a severe urinary tract infection that necessitated hospitalization and treatment with antibiotics; in patient 5, there was a dramatic improvement in serum bilirubin from 126–25 μM and an increase in serum albumin from 20–25 g/l. The levels of serum albumin and bilirubin for the individual patients are shown in Figure 5A–5E. Figure 5F and 5G shows the percentage changes in serum bilirubin and albumin levels for the entire group of five patients, and Figure 5H shows the disappearance of ascites as demonstrated by CT scan in patient 5.

Figure Figure 5..

Post-transplant follow-up of patients in phase I trial. (A–E): Changes in serum bilirubin and albumin levels after cell infusion in patients 1 to 5 (normal ranges: albumin 33–47 g/l, bilirubin 3–21 μM). (F): Percentage changes in serum bilirubin levels after cell infusion in patients 1 to 5. Sustained reductions in bilirubin were observed in three patients. (G): Percentage changes in serum albumin levels after cell infusion in patients 1 to 5. Increased levels of albumin were seen in four patients. (H): Computed tomography scans of the liver of patient 5 before and after cell infusion, showing resolution of the ascites 2 months after transplant. Abbreviation: UTI, urinary tract infection.


Cellular therapies based on stem cells and their derivatives are destined to revolutionize the practice of medicine in the near future. They hold the promise of cures for a great variety of conditions and diseases of tissue injury and degeneration ranging from Alzheimer's disease to trauma-induced injuries of the musculoskeletal system [17]. The essential requirements for stem cell therapy are (a) an easily procurable source of the stem cells themselves, (b) identification and characterization of the stem cell properties, (c) ability to increase (“expand”) cell numbers in culture reliably and reproducibly, (d) potential for differentiation of stem cell progeny into the desired tissue type, and (e) demonstration that the transplanted cells improve the function of damaged tissue. Other requirements that have been proposed are clonality and robustness. However, even cloned cell populations develop heterogeneity as they proliferate [18, 19]. It has been suggested, therefore, that cell population studies may be more important than clonality studies in defining the plasticity of stem cells, because plasticity may reflect the probability of stem cells differentiating into the appropriate lineage when they are placed in the correct microenvironment [9].

Adult human bone marrow and peripheral blood are easily available sources of stem cells. They contain two major types of stem cells, the hematopoietic stem cells and the MSCs. Classically, the hematopoietic stem cells are the source of all of the circulating mature blood cells, whereas the MSCs provide the stromal cells constituting the microenvironment within the marrow cavities. More recent data indicate broader potential for MSCs [20] in particular. Herein, we describe evidence that CD34+ cells have the ability to improve liver function in patients with liver disease.

Hematopoietic stem cells have been known to exist since the early studies of Till and McCulloch [21], whereas the MSCs were originally found in mouse bone marrow by Friedenstein et al. in 1974 [4] and later became known as bone marrow fibroblasts or stromal cells (reviewed in [22]). The term “mesenchymal stem cells” has been used only relatively recently. It has been known for a long time that cultures of bone marrow stromal cells may be induced to form adipocytes, osteoblasts, and endothelial cells [22] and that their phenotype includes expression of smooth muscle actin [23]. However, the phenotype of MSCs remains relatively poorly characterized [24] so that pure populations cannot be prospectively isolated from bone marrow. Similarly, the multipotential adult stem cells (MAPCs) [5] and USSCs [6] cannot be defined in freshly harvested hematopoietic tissue but are only characterized after prolonged in vitro culture [9]. In contrast, adherent CD34+ cells can be prospectively isolated from bone marrow or blood in known numbers and as a morphologically and immunophenotypically defined cell population. We acknowledge, however, that even highly purified stem cells are heterogeneous with regard to their functional characteristics and that they may exhibit phenotypic changes during the cell cycle [9]. It is relevant, therefore, that G-CSF-mobilized CD34+ cells have been demonstrated to be predominantly quiescent [25, 26], a feature that might be expected to contribute to their relative uniformity at the time of isolation.

The numbers of stem cells available to start a culture is a limiting factor in the progress of tissue regeneration from stem cells. Conveniently, there are large numbers of adherent CD34+ cells in “mobilized” PBPC (peripheral blood progenitor cell) harvests. Mobilization refers to the procedure whereby stem cells in the marrow are induced to enter the bloodstream when donors are treated with G-CSF and the circulating cells are harvested by leukapheresis. Typical yields of cells, obtained in our institution from donors for hematology patients, range from 5–10 × 1010, of which most are MNCs and approximately 1% are CD34+ (5–10 × 108). The CD34+ cells are separated using the CliniMACS device (a scaled-up version of the MiniMACS for clinical use). The adherent CD34+ cells constitute approximately 1% of the total CD34+ population so that a typical leukapheresis provides 5–10 × 106 putative stem/progenitor cells, and they have been successfully isolated in 100% of cases tested. Yields of this magnitude reduce the degree of cell number expansion and the time required to generate a clinically useful product.

In vitro culture of the adherent CD34+ cell fraction indicates the option to transplant their more differentiated progeny, which may be appropriate for some clinical applications. A 3- to 4-log expansion in cell number is achievable within 1–2 weeks (Fig. 3) and would provide 5–10 × 109 cells for clinical application. In our hands, the cells have been isolated in 100% of the cases tested. The cell expansion is also highly reliable and reproducible and has been successful in 98.4% of the 125 cultures we have initiated.

Both before and after culture, the adherent CD34+ cells and their progeny express an array of gene products as revealed by RT-PCR analysis, flow cytometry, and immunocytochemistry and adopt morphologies consistent with the tissue lineages shown in Figure 3. The RT-PCR studies are particularly crucial because it has been shown that some proteins, like albumin and insulin, can be taken up by the cells from the culture medium and are detectable by immunocytochemistry and flow cytometry. The RT-PCR results indicate that adherent CD34+ cells differentiate into cells expressing markers associated with, but not exclusive to, hematopoietic, hepatic, pancreatic, cardiovascular, and nervous tissues.

The phase I clinical study demonstrated the safety of administering G-CSF followed by leukapheresis and reinfusion of CD34+ cells in patients with liver insufficiency. It is important to note that the patients could respond to G-CSF treatment and that their white blood cell counts were increased in all cases, because this is a prerequisite for the remainder of the treatment protocol. However, the cell yields were lower than those expected from donors for hematology patients. Clinically, the procedure was well tolerated with no observed procedure-related complications. There were no cases of hepatorenal syndrome, and injection of the CD34+ cells into the hepatic artery or portal vein did not result in any thrombotic episode or bleeding after the percutaneous procedure. Importantly, there was some evidence of improvement in albumin and bilirubin levels, even though the trial was designed to be a safety and efficacy study.

Administration of G-CSF to liver failure patients being treated with interferon-γ did not result in any additional clinical improvement (N. Habib, unpublished data). Thus, whereas G-CSF-mobilized cells circulate through the liver, we propose that the relevant stem cells are only a tiny minority of the total population. Therefore, we speculate that stem cell engraftment in damaged liver is considerably enhanced by high concentrations of relevant stem cells delivered directly to the injured tissue. The distinct cellular and molecular properties of the adherent CD34+ cells lead us to believe that they, rather than nonadherent CD34+ cells, are responsible for the potential clinical benefit. At this stage, the mechanism for the effect on liver function is not clear but may reflect activation of genes corresponding to a hepatocyte differentiation program upon exposure to the injured liver environment.


Overall, these results are encouraging for the future development of stem cell therapy for patients with liver insufficiency. Currently, we are initiating a phase I/II clinical trial to test the hypothesis that cells derived from adherent CD34+ putative stem/progenitor cells are responsible for the clinical improvement seen in patients with liver damage.


M.Y.G., N.A.H., and J.P.N. own stock in LiverCyte Limited.


We thank Dr. A. Marrone for assistance with the TRAP assay, all the medical and nursing staff who were involved in the care of the patients in the phase I trial, and A. Deisseroth and Dr. A. Garen for their scientific advice. This work is based on international patent application PCT/GB2004/005365.