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Prepublished online as a Blood First Edition Paper on June 14, 2002; DOI 10.1182/blood-2002-02-0605.
GENE THERAPY
From Oxford BioMedica (UK) Ltd; Molecular Immunology
Unit, Institute of Child Health, London; Department of Molecular
Genetics, Institute of Ophthalmology, University College London; Renal
Section, Imperial College of Science Technology and Medicine, London,
United Kingdom.
Anemia is a common clinical problem, and there is much interest in
its role in promoting left ventricular hypertrophy through increasing
cardiac workload. Normally, red blood cell production is adjusted
through the regulation of erythropoietin (Epo) production by the
kidney. One important cause of anemia is relative deficiency of Epo,
which occurs in most types of renal disease. Clinically, this can be
corrected by supplementation with recombinant Epo. Here we describe an
oxygen-regulated gene therapy approach to treating homozygous
erythropoietin-SV40 T antigen (Epo-TAgh) mice with relative
erythropoietin deficiency. We used vectors in which murine Epo
expression was directed by an Oxford Biomedica hypoxia response element
(OBHRE) or a constitutive cytomegalovirus (CMV) promoter. Both
corrected anemia, but CMV-Epo-treated mice acquired fatal
polycythemia. In contrast, OBHRE-Epo corrected the hematocrit level in
anemic mice to a normal physiologic level that stabilized without
resulting in polycythemia. Importantly, the OBHRE-Epo vector had no
significant effect on the hematocrit of control mice. Homozygous
Epo-TAgh mice display cardiac hypertrophy, a common
adaptive response in patients with chronic anemia. In the
OBHRE-Epo-treated Epo-TAgh mice, we observed a significant
reversal of cardiac hypertrophy. We conclude that the OBHRE promoter
gives rise to physiologically regulated Epo secretion such that the
hematocrit level is corrected to healthy in anemic Epo-TAgh
mice. This establishes that a hypoxia regulatory mechanism similar to
the natural mechanism can be achieved, and it makes EPO
gene therapy more attractive and safer in clinical settings. We
envisage that this control system will allow regulated delivery of
therapeutic gene products in other ischemic settings.
(Blood. 2002;100:2406-2413) An important cause of anemia is relative deficiency
in the production of the glycoprotein hormone erythropoietin (Epo),
which regulates the formation of red blood cells (RBCs). Relative Epo deficiency occurs in almost all patients with chronic renal
failure.1,2 Decreased RBC production reduces the
oxygen-carrying capacity of the blood, resulting in tissue hypoxia.
Pathophysiologic consequences correlate with the severity of the anemia
and range from fatigue and reduced exercise tolerance to cardiac
hypertrophy. Erythropoietin deficiency in renal disease can be treated
remarkably effectively by regular administration of recombinant human
Epo (rhEpo) several times a week. Erythropoietin can also be used to
treat anemia in patients with cancer and chronic inflammatory diseases
such as rheumatoid arthritis.3,4 Although it is safe,
treatment with erythropoietin is relatively costly and entails some
inconvenience in monitoring and administration. Hence, there has been
considerable interest in developing a gene therapy strategy for the
delivery of Epo whereby single administration of the EPO
gene would ensure the long-term delivery of Epo.
Although the EPO gene has been delivered to animals using
various plasmid and viral vectors, the relevance of these studies to
chronic anemia and its sequelae have been limited. This is because the
EPO gene has been delivered to healthy
animals5-10 or has been used in models such as
We wanted to model a potential human clinical protocol, and it was
important that EPO gene therapy could be regulated.
Uncontrolled delivery of the EPO gene leads to an expanded
red cell mass (polycythemia),5,16 with hemodynamic and
rheologic problems including increased risk for vascular
thrombosis.17 One approach to control gene expression is
to use a regulated promoter that can switch EPO gene
expression on and off, such as the tetracycline- or
rapamycin-responsive promoters. To date, these promoters have only been
shown to regulate the hematocrit level above normal baseline rather
than to maintain normal levels.8,18-20 We envisage that
the use of these extrinsic regulation systems in a patient could be
costly and cumbersome, especially because of the addition of the
pharmacologic regulatory agents may interfere with other patient
medications. Therefore, we set out to develop a homeostatic system of
gene therapy based on sensing and correcting tissue hypoxia given the
normal regulation of the erythropoietin gene.
Reduced oxygenation of blood reaching the kidney is the key physiologic
signal for increasing erythropoiesis through increased expression of
the EPO gene by the fibroblasts of the renal cortex and
outer medulla14,21 and, to a lesser degree, in the adult by
the hepatocytes and Ito cells of the liver. In hypoxia, a heterodimeric transcription complex (hypoxia-inducible factor-1 [HIF-1]) binds to a
hypoxia-response element (HRE) lying 3' to the EPO gene to stimulate transcription.22 Although erythropoietin gene
expression is highly cell-type specific, most (if not all) cell types
activate the HIF-1 pathway in response to hypoxia.23,24
Consistent with this, many genes active in a broad range of cell types
and tissues are now known to contain HREs and to respond to
stabilization of the HIF-1 transcription factor in
hypoxia.25 Various natural and synthetic HRE-containing
promoters have been used to direct heterologous gene expression in
response to hypoxia Healthy and anemic mice
Cell lines
Transient transfections Typically, cells seeded in a 24-well dish were brought to 70% confluence and were transfected with 0.21 µg plasmid using the Fugene-6 transfection reagent (Boehringer Mannheim, Indianapolis, IN).Hypoxia in vitro Twenty-four hours after transduction or transfection, cells were either incubated for another 16 hours under normoxic conditions in a standard incubator (21% O2, 5% CO2, 74% N2) or under hypoxic conditions (0.1% O2, 5% CO2, 95% N2) using a multigas incubator purchased from Heto-Holten (Allerod, Denmark).Construction of recombinant AAV vectors The murine erythropoietin cDNA was cloned through nested polymerase chain reaction (PCR) on murine kidney cDNA (Quickclone cDNA; Clontech, United Kingdom) using 2 pairs of nested PCR primers: primer set 1, 5'-GACAGTGACCACTTTCTTCCAG-3' and 5'-GGACAGACTGGTAAGAAGGTAATG-3'; primer set 2, 5'-CAGCTAGGCGCGGAGATG-3' and 5'-CAGCAGCATGTCACCTGTC-3'.The mEpo PCR product was cloned into the pUC18 plasmid (Panvera, Madison, WI) and was subsequently removed as an XbaI-EcoRI fragment and cloned in to the pCI-Neo (Promega, Southampton, United Kingdom) NheI-EcoRI sites to create pCMV-Epo. The CMV/IE promoter in pCMV-Epo was replaced with the OBHRE promoter to create pHRE-Epo.27 An oligonucleotide was cloned into the BamHI and SpeI restriction sites in the multiple cloning site of the pSL1180 plasmid (Amersham Pharmacia Biotech, Buckinghamshire, United Kingdom) to generate the following restriction sites: BamHI-NheI-MluI-XhoI-StuI-NruI-BclI-SpeI-BglII. The AAV-CMVEpo vector genome was constructed by creating a 145-base
pair (bp) oligonucleotide consisting of the wild-type AAV-2 inverted
terminal repeat (ITR) (GenBank accession number: NC_001401) flanked by
BamHI and NheI compatible ends. The ITR was
cloned sequentially in reverse and forward orientation into the
BamHI-NheI, SpeI, and BglII
sites of the modified pSL1180 vector. The CMV-Epo
BsaBI-BglII fragment from pCMVEpo was cloned into
the StuI-BglII sites of the modified pSL1180
vector, together with a 1.7-kb BclI-BglI stuffer
fragment from the LacZ gene such that the complete internal
cassette measured 4.2 kilobases (kb). The AAV-HREEpo vector genome was
created by exchanging the CMV/IE NotI- Eco47III
promoter fragment in AAV-CMVEpo for the OBHRE
NotI-XmnI promoter fragment in pHRE-Epo (Figure
1A).
Recombinant AAV-2 vectors were produced according to the published method.31 AAV particles were determined by dot blot quantification of genome copy and direct comparison with a recombinant AAV vector expressing CMV-GFP of known biologic titer. Spleen cell proliferation assay The functionality and regulation of the cloned Epo cDNA was verified using a biologic spleen cell proliferation assay based on a published method.32 Briefly, 2- to 3-month-old mice (C57BL/6J × C3H/HeB F1 hybrid), each weighing 25 to 35 g, were given 2 consecutive daily intraperitoneal injections of 60 mg/kg phenylhydrazine hydrochloride. Spleens were isolated 3 days after the second injection. Single-cell suspensions from the spleen were prepared 3 days after the second injection and were seeded into black-walled, 96-well plates (Canberra Packard, Mississauga, ON, Canada) at a density of 4 × 105 cells per well. Supernatants were collected from HT1080 cells 5 days after transfection with either pCMV-EPO or pHRE-EPO plasmids, and 1 µL of each supernatant was added to the splenocyte cell cultures. As a positive control, rhEpo was used at 500 U/mL. Splenocyte cell cultures were incubated for 22 hours and then were assayed for proliferation using a chemiluminescent 5-bromodeoxyuridine (BrdU) assay (Roche, Mannheim, Germany).Detection of erythropoietin in vitro Erythropoietin was detected in cell supernatants using the Quantikine IVD Epo enzyme-linked immunosorbent assay (ELISA) kit at a detectable threshold of 2 mU/mL (R&D Systems, Abingdon, Oxon, United Kingdom).Histologic analyses Standard hematoxylin and eosin staining was carried out for assessment of tissue morphology. To determine the distribution of iron in the liver, sections were stained using Perl Prussian blue. For immunologic analysis, tissue sections were fixed in acetone. The TER-119 antibody (BD PharMingen, Oxford, United Kingdom) was used to recognize cells committed to the erythroid lineage, from proerythroblasts to mature erythrocytes. It was used at a dilution of 1:50 overnight followed by the horseradish peroxide (HRP)-conjugated secondary antibody (HRP rabbit anti-rat; DAKO, Glostrup, Denmark) at a dilution of 1:50 in 10% mouse serum. DAB (3,3'-diaminobenzidine) was added for 10 minutes, and the slides were counterstained with hematoxylin.For electron microscopy, the heart was isolated from one mouse from each of the untreated and the AAV-HREEpo-treated healthy and Epo-TAgh groups on day 189 of the study. The hearts were dissected into 1-mm cubes and were immersion-fixed in 1% gluteraldehyde/2.5% paraformaldehyde. Samples were washed in phosphate-buffered saline (PBS) and were postfixed in 1% OsO4 in 0.1 M phosphate buffer, washed in distilled water overnight at 4°C, dehydrated in alcohol, and embedded in Durcupan epoxy resin. Ultrathin cross-sections of the myocardium were stained with 2% uranyl acetate, followed by 1% lead citrate (Reynold stain) and were examined under the Philips 401 transmission electron microscope (Wilmington, MA). Sarcomere measurements were made from randomly taken photographs. Statistical analysis The unpaired Student t test was used to determine whether there was a significant difference between groups of data. Groups were considered significantly different when P < .05.
Hypoxia-mediated regulation of functional murine Epo expression in vitro We have observed that a synthetic HRE multimer, referred to as OBHRE, can combine a good induction ratio with a high level of expression comparable to that achieved by strong constitutive promoters such as the CMV promoter, but only when the oxygen concentration is low.27 The OBHRE promoter was inserted into plasmid and AAV vectors to produce pHRE and AAV-HRE, respectively (Figure 1A). Similar vectors containing the human CMV promoter were pCMV and AAV-CMV. A cDNA for murine EPO was inserted into these vectors, and green fluorescence protein (GFP)-expressing vectors were used as negative controls. We wanted to use the murine EPO rather than the human EPO to avoid immune responses that would compromise the efficacy of the gene therapy. We first confirmed that the murine EPO gene functioned in vitro. The production of mEpo in the culture supernatant of HT1080 cells, transfected with pHRE-Epo or pCMV-Epo and maintained in normoxia or hypoxia, was determined using a spleen cell proliferation assay (Figure 1B). Both plasmids directed the expression of functional mEpo, but with pHRE-Epo the expression was at least 8-fold higher from cells maintained in hypoxia compared with normoxia. Similarly, the recombinant AAV vectors were transduced into T47D cells, placed in normoxia, or exposed to hypoxia for 16 hours and then returned to normoxia (Figure 1C). Secretion of mEpo into the supernatant was assessed in an Epo ELISA 1 day and 4 days after hypoxic induction. AAV-CMV-directed mEpo expression increased during the 4 days in normoxia and hypoxia, whereas AAV-HRE-directed mEpo expression increased from basal levels only in the hypoxia-exposed cultures as measured at day 1. By day 4, levels of mEpo had returned to baseline. Collectively, these data from the 2 assays indicated that the mEPO gene was functional and that the expression could be activated by hypoxia and switched off in normoxia. This reversible expression was the profile that would be required for a gene therapy vector that could deliver Epo under anemic conditions but that would be shut down once normal oxygenation was restored.Regulated delivery of Epo in vivo In preliminary studies we examined the Epo-TAgh hindlimb skeletal muscle for evidence of hypoxia. As expected, immunologic analysis for CD31 showed a modest increase in muscle capillarity. Although variable, immunolabeling for vascular endothelial growth factor (VEGF) showed more signal in specimens from the Epo-TAgh mice than control mice (data not shown). This suggested that the skeletal muscle was likely to be sufficiently hypoxic to activate the OBHRE promoter because VEGF gene expression is activated by hypoxia, predominantly through the HIF-1-mediated transcriptional pathway.Healthy or Epo-TAgh mice were injected in the left hindlimb
with AAV-CMVGFP-, AAV-CMVEpo-, or AAV-HREEpo-recombinant AAV
viruses. Hematocrit measurements were made regularly over a period of 7 months (Figure 2). The control vector was
AAV-CMVGFP, and this produced no change in the hematocrit level of
healthy mice, which was maintained at approximately 52% (Figure 2A,
open squares), or of Epo-TAgh mice, which was maintained at
approximately 18% (Figure 2A, closed squares) throughout the duration
of the study. These levels were identical to those of untreated
controls (Figure 2A, healthy mice, open circles; Epo-TAgh
mice, closed circles). In marked contrast, when the healthy and Epo-TAgh mice were injected with the constitutive Epo
vector, AAV-CMVEpo, there was a dramatic increase in the hematocrit
level in each group that was significant after only 7 days and that
increased to 85% after 45 days (Figure 2A, diamonds). Two mice in this
group died suddenly at day 60, by which time the blood in the remaining animals became too viscous to obtain samples for hematocrit analysis; therefore, the animals in these groups were killed. However, a different result was obtained when the hypoxia-regulated vector, AAV-HREEpo, was used. In healthy mice (Figure 2, open triangles) there
was no statistically significant effect on the hematocrit level after
AAV-HREEpo treatment. In the Epo-TAgh mice, the AAV-HREEpo
vector led to a steady increase in the hematocrit of 2% to 2.5% per
week for 11 weeks, until a plateau was reached at the normal level.
This plateau was an average hematocrit level between 46% and
50%
Stimulation of extramedullary hematopoiesis in AAV-HREEpo-treated Epo-TAgh mice To further characterize the response to regulated erythropoietin in Epo-TAgh mice, we analyzed the liver, spleen, and bone marrow for evidence of hematopoiesis. For this we used TER-119, an antibody that recognizes cells in the late stages of the erythroid lineage.33 The liver and bone marrow showed no discernible difference in the TER-119 staining in all groups examined. However, the spleen taken from the Epo-TAgh mice treated with AAV-HREEpo (Figure 3D) contained elevated levels of erythroid (TER-119+) cells in the red pulp region compared to the spleens taken from untreated Epo-TAgh (Figure 3B) mice. Spleens taken from untreated and AAV-HREEpo-treated healthy mice did not show any difference in the level of erythroid cells (Figure 3A,C). This indicates that the production of Epo from the AAV-HREEpo vector stimulated extramedullary hematopoiesis in the spleens of the Epo-TAgh mice but not in the healthy mice.
Impact of AAV-HREEpo on liver iron loading in Epo-TAgh mice It has previously been shown that the Epo-TAgh mice display hepatic iron loading because of enhanced mucosal iron uptake in the absence of erythropoiesis.34 Hepatic iron loading can lead to functional impairment in the liver. We were interested to see the distribution of nonheme iron in the livers of untreated and AAV-HREEpo-treated Epo-TAgh mice using Perl Prussian blue. Livers from untreated Epo-TAgh mice (Figure 3F) showed an increase in the amount of nonheme iron in the cytoplasm of the liver Kupffer and parenchymal cells compared with the livers from healthy mice (Figure 3E). Interestingly, the livers from AAV-HREEpo-treated Epo-TAgh mice showed a decrease in the amount of nonheme iron. This suggests the stored iron has been mobilized from the liver and used for erythropoiesis.Organ analysis of study animals We wanted to determine whether Epo gene therapy caused any further structural changes to internal organs. Changes in red blood cell composition affect the volume and the pressure of blood. In chronic anemia, this hemodynamic alteration leads to gradual development of cardiac enlargement (hypertrophy) as the cardiac output increases to compensate for the decreased oxygen-carrying capacity of the blood. We compared the weights of some of the organs in the untreated and treated Epo-TAgh and healthy mice at the end of the experiment (Figure 4). There was no significant difference between any of the groups in brain size. However, a marked difference was noted in the spleen. The AAV-CMVEpo-treated healthy and Epo-TAgh mice had significant splenomegaly. This is most likely a result of extramedullary hematopoiesis and vascular congestion caused by the increase in RBC load. Consistent with this, AAV-HREEpo-treated Epo-TAgh mice had only slightly larger spleens than their healthy counterparts.
Hearts taken from the untreated Epo-TAgh mice weighed 200% more than those taken from healthy mice, consistent with anemia-associated hypertrophy. Overproduction of Epo from the AAV-CMVEpo vectors had no significant effect on heart weight in either the Epo-TAgh or the healthy mice. Most interesting was the reduction in heart weight in the AAV-HREEpo-treated Epo-TAgh mice such that it was now only 33% greater than in healthy mice, showing that the hypertrophy had partially been reversed. AAV-HREEpo treatment had no effect on heart weight in the healthy mice. Histologic and ultrastructural analysis of the hearts confirmed gross
hypertrophy in the Epo-TAgh mice (Figure
5). Hypertrophy occurs as a result of an
increase in the size of individual cells, and this can be seen in the
Epo-TAgh heart as an increase in the average length of the
sarcomere (the width between the muscle striations) from 0.97 ± 0.12
µm in healthy mice to 1.20 ± 0.20 µm in Epo-TAgh
mice. AAV-HREEpo treatment had no effect on sarcomere length in the
healthy mice; however, in the Epo-TAgh mice, it was reduced
to 1.05 ± 0.08 µm, further confirming partial correction of the
cardiac hypertrophy in the AAV-HREEpo-treated Epo-TAgh
mice.
This study demonstrates physiologically controlled regulation of Epo gene expression in genetically anemic Epo-TAgh mice. Each mouse received a single intramuscular dose of recombinant AAV vector, where EPO expression is regulated by the OBHRE promoter, AAV-HREEpo. The Epo-TAgh mice treated with AAV-HREEpo showed correction of the hematocrit to a physiologically normal level that was stabilized for the duration of the 7-month experiment. In other studies, sequences from the native EPO enhancer have been used to boost expression of EPO in an adenovirus or in encapsulated skeletal muscle cells.35,36 However, this study is the first example of the physiological regulation of EPO expression using an optimized hypoxia-responsive promoter in a clinically relevant anemic animal model. These data illustrate that the OBHRE promoter can sense the level of tissue hypoxia in the skeletal muscle and switch the expression of EPO on and off accordingly. The physiological regulation of the AAV-HREEpo vector in the skeletal muscle was further reinforced by the finding that it had no effect on the hematocrit level in healthy mice. It is perhaps surprising that such exquisite physiological regulation was achieved in the skeletal muscle because it has been argued that the kidney is uniquely suited to sensing the hematocrit and regulating the production of Epo.37 These data suggest that the ability to sense and respond effectively to changes in the hematocrit may not be limited to the kidney and liver. The correction of chronic anemia is a clinically important
challenge. Anemia alone can lead to cardiac hypertrophy in the absence
of any other cardiovascular disorder, and in renal patients anemia is
an important contributor to left ventricular hypertrophy. Importantly,
in patients with renal disease, the degree of left ventricular
hypertrophy is a strong independent predictor of death.38 In the presence of heart disease, especially coronary artery disease, anemia intensifies angina and contributes to a high incidence of
cardiovascular complications. Clinically, it is essential to tailor the
dose of rhEpo carefully to obtain an optimal response and to avoid
complications related to polycythemia. The appropriate dose not only
varies from patient to patient, it requires frequent adjustment based
on hematocrit level. The approach we have taken here has 3 main
advantages. First, the results described in this study suggest that an
EPO therapy regulated by the OBHRE promoter might require
less tailoring to individual patient requirements. Second,
EPO gene therapy might achieve more precise correction of
anemia, allowing more normal functioning of important organs such as
the heart. The potential of this is illustrated by the substantial
correction of left ventricular hypertrophy in the Epo-TAgh
mice in this study. Third, EPO gene therapy might be
used in conditions such as myeloma, cancer, and thalassemia,
which are often relatively resistant to Epo and would require higher
doses than those administered to renal patients. Illustrating the
problem of unregulated erythropoietin in this setting, when
Epo-secreting hematopoietic cells were engrafted into a mouse model of
We have clearly shown in this study that the OBHRE promoter
functions to gives rise to physiologically controlled regulation of
EPO gene expression in a relevant anemic mouse model. Before we evaluate this gene therapy approach clinically, we would like to
test the activity of the OBHRE promoter in larger animal models of
Epo-responsive anemia Although recombinant Epo has good safety and clinical records, the exquisite regulation observed in this model may lead to the consideration of genetic delivery strategies. In addition, the system described here shows the potential of combining efficient gene transfer to the skeletal muscle, with the individual's own cells supplying the regulatory component of the transcriptional machinery, HIF-1. We envisage that this approach could be useful in controlled local delivery of other polypeptides in diverse ischemic diseases. In summary, this study describes the development of a physiologically regulated gene therapy modality that can deliver long-term, physiologically regulated expression of EPO, leading to correction of the hematocrit level and associated cardiac hypertrophy in chronically anemic mice.
We thank Peter Ratcliffe (The Henry Wellcome Building of Genomic Medicine, Oxford, United Kingdom) for helpful contributions to the work and Michael Dennis (CAMR, Salisbury, Wiltshire, United Kingdom) for maintenance and management of the animals. We also thank Sarah Busby (Department of Pharmacology, University of Oxford, United Kingdom) for the electron microscopy work.
Submitted March 14, 2002; accepted May 14, 2002.
Prepublished online as Blood First Edition Paper, June 14, 2002; DOI 10.1182/blood-2002-02-0605.
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
Reprints: Katie Binley, Oxford BioMedica (UK) Ltd, Medawar Centre, Oxford Science Park, Oxford OX4 4GA, United Kingdom; e-mail: k.binley{at}oxfordbiomedica.co.uk.
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Abstract
Introduction
-thalassemic mice
for example, tumor cells, muscle cells,
and macrophages.
307/
represents Epo-TAgh mice;
, healthy mice.
