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. 2009 Apr 14;17(7):1155–1163. doi: 10.1038/mt.2009.65

Short-term Correction of Arginase Deficiency in a Neonatal Murine Model With a Helper-dependent Adenoviral Vector

Chia-Ling Gau 1, Robin A Rosenblatt 2, Vincenzo Cerullo 3,4,5, Fides D Lay 2, Adrienne C Dow 2, Justin Livesay 6, Nicola Brunetti-Pierri 3,4, Brendan Lee 3,4, Stephen D Cederbaum 1,7,8, Wayne W Grody 1,6,7, Gerald S Lipshutz 2
PMCID: PMC2835205  PMID: 19367256

Abstract

Neonatal gene therapy has the potential to ameliorate abnormalities before disease onset. Our gene knockout of arginase I (AI) deficiency is characterized by increasing hyperammonemia, neurological deterioration, and early death. We constructed a helper-dependent adenoviral vector (HDV) carrying AI and examined for correction of this defect. Neonates were administered 5 × 109 viral particles/g and analyzed for survival, arginase activity, and ammonia and amino acids levels. The life expectancy of arg−/− mice increased to 27 days while controls died at 14 days with hyperammonemia and in extremis. Death correlated with a decrease in viral DNA/RNA per cell as liver mass increased. Arginase assays demonstrated that vector-injected hepatocytes had ~20% activity of heterozygotes at 2 weeks of age. Hepatic arginine and ornithine in treated mice were similar to those of saline-injected heterozygotes at 2 weeks, whereas ammonia was normal. By 26 days, arginase activity in the treated arg−/− livers declined to <10%, and arginine and ornithine increased. Ammonia levels began increasing by day 25, suggesting the cause of death to be similar to that of uninjected arg−/− mice, albeit at a later time. These studies demonstrate that the AI deficient newborn mouse can be temporarily corrected and rescued using a HDV.

Introduction

Arginase deficiency is a rare metabolic disorder resulting from a loss of arginase I (AI), the sixth and final enzyme in the urea cycle, which is the major pathway for the detoxification of ammonia in mammals. AI is expressed most prevalently in the liver, where in coordination with the other enzymes of the urea cycle, sequestration of nitrogen as urea occurs.1 AI hydrolyzes arginine into ornithine, which can re-enter the urea cycle, and urea that gets excreted as waste. Arginase deficiency is the least severe of the urea cycle disorders, characterized by hyperargininemia and infrequent episodes of hyperammonemia. Human patients suffer from neurological impairment with cortical and pyramidal tract deterioration, spasticity, loss of ambulation, seizures, and severe mental and growth retardation. However, they usually avoid the catastrophic hyperammonemic crises characteristic of the other urea cycle disorders and, therefore, tend to survive much longer.1 The lack of frequent episodes of hyperammonemia is probably due to an increase in the second form of arginase, AII, which compensates for the lack of AI.1 This enzyme is expressed in extrahepatic tissues, mainly in the kidney and prostate.1,2 Although dietary and pharmaceutical interventions can partially alleviate AI deficiency, there is no completely effective therapy available today.

AI-deficient C57Bl/6 mice were previously generated in our laboratory by replacing exon 4, the active site of the AI gene with the neomycin resistance gene.3 No AI RNA was found on northern blots nor was any crossreacting material detected. These mice completely lack liver AI activity. In contrast to the human disease in which patients can survive into adulthood, C57Bl/6 mice with AI deficiency die between postnatal days 10–14 due to severe hyperammonemia.1,3 Plasma ammonia levels of AI-deficient mice in metabolic crisis are increased greater than tenfold and their livers are abnormal with histopathologic features similar to those seen in human arginase-deficient patients who died with hyperammonemia later in life.3

Adenoviral vectors can transduce both dividing and postmitotic cells of nearly all tissues including skin, muscle, arteries, bone, nerve, and liver.4 Helper-dependent adenoviral vectors (HDVs) lack all viral coding sequences, leaving only the inverted terminal repeats necessary for vector propagation and the ψ sequence required for packaging. In general, they result in long-term transgene expression without chronic toxicity.5,6,7,8,9 This is in contrast to first generation adenoviral vectors, where adaptive immune responses led to a loss of expression and hepatic transaminitis. These features of HDV suggest that they may be well-suited and an ideal choice for vector-mediated hepatic gene therapy for metabolic diseases.

Early postnatal gene therapy has the potential to ameliorate genetic abnormalities before the development of phenotypic disease. In adults, rapid cellular proliferation is uncommon. Individual hepatocytes in the adult mouse liver are replaced once every 180–400 days.10,11 Conversely, in the neonatal murine liver, the rate of hepatocellular proliferation is much higher and cellular proliferation may affect episomal vector genomes.12,13 We evaluated the efficacy of HDV-mediated gene transfer to correct the biochemical defect in the neonatal murine model of AI deficiency. We assessed the survival of animals after a single neonatal intravenous dose of HDV expressing AI. Furthermore, we assessed the sites of tissue expression, transgene expression, and arginase expression in the liver and determined the growth in mass of liver in relation to the growth of the mouse in the first 7 weeks of life. Finally, we examined plasma ammonia levels and amino acid profiles. Long-term correction of the deficiency, if attained, would allow for examination of the effect of AI deficiency on the brain and for study of the role of excess arginine in neurodegeneration.

Results

The lifespan of arginase-deficient mice is prolonged with HDV-mAI

One of the mysteries of arginase deficiency has been the role of arginine in neurodegeneration. Because AI-deficient mice die by postnatal day 14 from liver failure with no visible signs of neurodegeneration, it was hypothesized that generating an adenoviral vector to express AI, specifically in liver, would rescue arginase-deficient mice from death. Extended survival of the mice would then allow for the examination of the long-term effect of arginase loss in the brain. Murine AI (mAI) was inserted downstream of the liver-specific PEPCK promoter and upstream of the woodchuck hepatitis virus post-transcriptional regulatory enhancer (WPRE), which was included to improve mRNA stability and, therefore, transgene levels (Figure 1). Heterozygote mAI-deficient mice were mated to produce litters containing wild-type, heterozygous, and homozygous-affected mice. All pups were injected intravenously through the superficial temporal vein between 2 and 3 days of age with either pharmaceutical grade saline or 5 × 109 viral particles/g of HDV-mAI and were genotyped 7 days after birth. Arg−/− mice injected with HDV-mAI (n = 15) survived up to 27 days as opposed to saline injected arg−/− mice (n = 15) that survived up to 14 days of life (Figure 2) (P <0.0001). In contrast to uninjected arg−/− mice, which exhibit a tremor phenotype at least 24 hours before death, HDV-mAI mice did not exhibit a tremor phenotype until <12 hours before death. No differences in phenotype were seen between mice injected on different days. Injection of HDV-mAI had no effect on the life span or phenotype of wild-type or heterozygous mAI mice (data not shown).

Figure 1.

Figure 1

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Schematic of HDV-mAI. The figure (not drawn to scale) shows the relevant portions of the helper-dependent adenoviral vectors containing (a) the mAI cDNA or (b) LacZ cDNA downstream of the liver specific PEPCK promoter and the ApoAI intron. ApoAI, ApoAI intron; HDV, helper-dependent adenoviral vector; hGH pA, human growth hormone polyA; ITR, inverted terminal repeat; LacZ, β-galactosidase; mAI, murine arginase I; WPRE, woodchuck postregulatory enhancer; ψ, packaging signal.

Figure 2.

Figure 2

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Survival of AI knockout mice. Arg+/− mice were mated to produce litters containing arg+/+, arg+/−, and arg−/− mAI mice. All pups were injected intravenously via the superficial temporal vein between 2 and 3 days of age with either pharmaceutical grade saline or 5 × 109 viral particles/g (5 × 1012 vp/kg) of HDV-mAI. AI knockout mice injected with HDV-mAI survived up to 27 days compared to saline-injected knockout mice, which survive up to 14 days postnatally. HDV, helper-dependent adenoviral vector.

Expression of HDV

To examine the biodistribution of the vector after a single neonatal injection of 5 × 109 viral particles/g helper-dependent adenovirus, we injected mice with HDV-LacZ, a β-galactosidase-expressing vector and examined the biodistribution of vector in various tissues 2 days later by quantitative real-time PCR (Figure 3a). As expected with a hepatotropic viral vector, the largest number of viral copies per cell was in the liver (10.4 ± 1.5 viral copies per cell). This was followed at substantially reduced copy number per cell by the spleen (0.16 ± 0.09 viral copies per cell), heart (0.15 ± 0.04 viral copies per cell), kidney (0.13 ± 0.07 viral copies per cell), lung (0.05 ± 0.02 viral copies per cell), muscle (0.011 ± 0.005 viral copies per cell), intestine (0.008 ± 0.003 viral copies per cell), brain (0.005 ± 0.003 viral copies per cell), and thymus (0.004 ± 0.001 viral copies per cell). Histochemical staining for β-galactosidase demonstrated patchy staining throughout the liver not localized to any particular portion of the liver lobule (Figure 3b).

Figure 3.

Figure 3

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Distribution of HDV after neonatal intravenous injection is widespread. (a) Biodistribution of helper-dependent adenoviral vector (HDV). Arg+/− mice were injected on the second day of life with 5 × 109 viral particles/g HDV-LacZ. Two days later, mice (n = 4) were euthanized and tissues were removed. Biodistribution of the hepatotropic HDV vector was determined by PCR. (b) Histochemical hepatic staining for helper-dependent adenoviral vector. X-gal staining of the liver was performed on the eighteenth day of life from mice that received a neonatal injection of HDV-LacZ. X-gal staining is detected in hepatocytes with no preference for any specific lobular location.

Both HDV-injected and saline-injected control mice were euthanized at preset time points. At each time point for removal for tissue analysis, mice were inspected macroscopically and no gross pathology was observed. Whole livers (n = 3–8 per time point) were removed and there were no tumors seen at all time points, ranging from 4 to 100 days. Total body weight and liver weight were recorded (Figure 4a,b). There were no differences in the animal weights or liver weights of animals that received HDV or saline. The liver generally ranged from 4.5% (on the second day of life) to 5.5% (at 7 weeks) of the animal's total weight. The average weight of the liver at injection on the second day of life was 0.0633 ± 0.0006 g, while the weight of the mouse was 1.34 ± 0.15 g. By 27 days of life, mice weigh 14.55 ± 1.05 g with a liver weight of 0.84 ± 0.11 g, a 10.9- and 13.3-fold increase, respectively. At 7 weeks, the liver weighed 1.12 ± 0.28 g and the weight of the mouse was 20.16 ± 3.76 g; some variation is present depending on mouse gender that becomes more pronounced as the mice become older. This represents an increase of 17.7-fold for the liver and 14.5-fold for the mouse over the first 7 weeks of life.

Figure 4.

Figure 4

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Growth of mouse and liver from neonate to adult. Both saline-injected and HDV-injected mice were euthanized at predetermined time points (n = 3–8 per time point). (a) Animals and (b) livers, after removal, were weighed. The mouse liver generally represents 4.5–5.5% of the weight of a lean C57BI/6 mouse (g = grams). HDV, helper-dependent adenoviral vector.

Viral copy number per hepatocyte (LacZ) was followed over time (Figure 5a). The number of copies of the replication-deficient virus per hepatocyte declined as the animal weight increased. Copies as high as 10.35 ± 1.46 per cell were found 2 days after injection, declining to 0.016 ± 0.003 copies per cell at day 40 of life. Although initially higher, RNA copies declined over time and by day 24 postnatally they reach their nadir. Viral mRNA declined from 745.0 ± 292.0 copies on the fourth day of life to 75.3 ± 11.1 copies on the fortieth day of life (Figure 5b).

Figure 5.

Figure 5

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Change in viral copies and message over time. (a) Viral copy number per cell was determined by quantitative real-time PCR, whereas (b) mRNA copies were similarly determined after reverse transcription.

To further examine viral RNA expression, HDV-mAI-injected wild-type and heterozygous mAI mice were euthanized at various time points to examine for the expression of viral mRNA (Figure 6a). The results demonstrate that viral RNA is present only in the liver, not in the brain.

Figure 6.

Figure 6

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Arginase expression by HDV-mAI decreases with time. (a) Tissue expression of viral RNA over time. Total RNA was isolated from tissues of HDV-mAI injected heterozygous mice at various ages (d = day). cDNA was synthesized and amplified using primers specific to the viral cDNA and analyzed by ethidium bromide-stained gel electrophoresis. The expected fragment sizes for HDV-mAI and β-actin cDNA are 750 and 350 bp, respectively; β-actin gDNA fragment size is 1 Kb. HDV-mAI cDNA was expressed only in the liver (1) and not in the brain (2). (b) Protein expression of HDV-mAI. Total protein was isolated from mice injected with HDV-mAI at postnatal days 14 or 25 (d = day). Western blot analysis demonstrated that arg−/− mice injected with HDV-mAI expressed subendogenous levels of arginase (arginase molecular size is 35 kd). Protein expression was greatly reduced when assayed on day 25. (c) Liver arginase activity of HDV-mAI-injected mice. Total protein was isolated from the livers of mice injected with either saline or HDV-mAI. Arginase activity was measured with a colorimetric assay determining the quantity of urea converted from arginine by each tissue lysate. Saline-injected arg−/− mice had virtually no arginase activity, whereas the livers of HDV-mAI-injected arg−/− mice at postnatal day 14 have ~20% of the arginase activity of heterozygous mice. By day 26, when the HDV-mAI-injected arg−/− mice have a short remaining lifespan, the hepatic arginase activity is <10% of endogenous activity. Error bars represent mean ± SD. HDV, helper-dependent adenoviral vector; KO, knockout; mAI, murine AI.

Injected arg−/− mice were euthanized at day 14 (n = 7) and 25 (n = 4) to examine protein expression of HDV-mAI in the liver. Western blot analysis demonstrated that arg−/− mice injected with HDV-mAI expressed subendogenous levels of mAI, when compared to heterozygous mice, and that protein expression was substantially decreased when assayed on day 25 (Figure 6b). A functional protein assay was performed by measuring arginase activity with a colorimetric assay determining the quantity of urea converted from arginine by each tissue lysate (Figure 6c). Saline-injected arg−/− mice (n = 6) have virtually no arginase activity in the liver (3.7 ± 1.3 nmol urea/µg protein/hour) at 14 days of life. At postnatal day 14, the liver arginase activity of HDV-mAI-injected arg−/− mice (n = 7) are increased due to the vector expressing AI (50.5 ± 35.4 nmol urea/µg protein/hour). Although this increase in arginase activity in vector-treated mice is not statistically significant at day 14, there is a trend toward statistical significance (P = 0.07). This is ~21.8% of the arginase activity of heterozygous mice (231.6 ± 68.8 nmol urea/µg protein/hour). By day 26, however, when the HDV-mAI-injected arg−/− mice (n = 9) are on the verge of death, liver arginase activity is <10% (21.0 ± 6.7 nmol urea/µg protein/hour) of endogenous activity of heterozygous mice, substantially less than at day 14.

The decline in vector-driven mAI expression is consistent with the timing of the death of the injected mAI knockout mice. As demonstrated, the viral copy number per cell declines early in the time course and by day 24 it reaches a relative steady-state but low level. ELISA and histopathological studies were performed to examine whether host humoral and cellular immune responses to the virus and transgene-encoded protein were present. No antiadenoviral antibodies nor difference from controls was detected (Figure 7a) from plasma samples at day 25 of life. No antiarginase antibodies were detected in arg−/− mice that received HDV-mAI on the second day of life and were tested at day 25 (Figure 7b). Histopathological evaluation of the liver demonstrated no lymphocytic infiltrates characteristic of a cytotoxic T lymphocyte–mediated response in either HDV-injected or saline control animals (Supplementary Figure S1).

Figure 7.

Figure 7

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Antibodies against adenoviral capsid proteins and against arginase. (a) Antiadenoviral-specific ELISA was performed on day 25 of mice injected with either saline or HDV-LacZ on the second day of life. There were no differences between the virus-injected or saline-injected groups. (b) Antiarginase-specific ELISA was performed on day 25 of arg−/− mice injected as neonates on the second day of life. There was no development of antiarginase antibodies (n ≥ 3 per group). Error bars represent mean ± SD. HDV, helper-dependent adenoviral vector; mAI, murine AI.

Rescue of metabolic phenotype

Arginase-deficient mice exhibit a series of metabolic changes. Most significantly, the C57Bl/6 knockout strain becomes hyperammonemic by postnatal days 12–14. This buildup of ammonia occurs due to the inadequate amount of arginase and the ability to eliminate ammonia as urea. Plasma ammonia levels were examined in mAI heterozygous and arg−/− mice, injected with either saline or HDV-mAI, at days 14 and 26 of life (Figure 8a). At day 14, saline-injected arg−/− mice (n = 10) were in extremis and exhibited hyperammonemia [684.6 ± 188.4 µmol/l ammonia versus 194.0 ± 59.4 µmol/l in saline-injected heterozygous mice (n = 12)] (P = 0.0001). HDV-mAI-injected arg−/− mice (n = 9) had ammonia levels similar to heterozygous mice (192.9 ± 69.2 µmol/l) (P = 0.821 when compared to heterozygous mice). At day 26, arg−/− mice injected with HDV-mAI (n = 7) have increased ammonia levels compared to heterozygous mice (n = 13) (387.7 ± 71.4 µmol/l versus 149.9 ± 38.2 µmol/l) (P = 0.0001), but not as high as saline-injected arg−/− mice at day 14 (P = 0.008). In addition, HDV-mAI injected arg−/−mice at day 26 did not exhibit a tremor phenotype in contrast to uninjected or saline-injected arg−/− mice at day 14, which exhibit a severe tremor phenotype.

Figure 8.

Figure 8

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Near correction of metabolic abnormalities occurs at day 14 of life in HDV-injected mice, but is lost by day 26. (a) Plasma ammonia of HDV-mAI injected mice. Plasma of mice injected with saline or HDV-mAI was collected at postnatal day 14 or 27 by cardiac puncture or retro-orbital bleeding (n = 9) for saline injected arg−/− mice at day 14, n = 6 for HDV-mAI injected arg−/− mice at day 14, n = 4 for HDV-mAI injected arg−/− mice at day 27, and n = 3 for arg+/− mice at each time point). At postnatal day 14, saline-injected arg−/− mice are on the verge of death and exhibit hyperammonemia, whereas arg−/− mice injected with HDV-mAI have levels of ammonia similar to those in uninjected heterozygous mice. By day 26, arg−/− mice that had received HDV-mAI, but which did not yet exhibit a tremor phenotype, had increased ammonia levels over heterozygous controls. Error bars represent mean ± SD. (b,c) Hepatic arginine and ornithine levels in HDV-mAI injected mice. Liver tissue was collected from HDV-mAI-injected mice at postnatal day 14 and 26 (n = 6 mice of each genotype at each time point). Arg−/− mice injected with HDV-mAI exhibited lower levels of arginine (b) and normalization of ornithine (c) at postnatal day 14 compared to saline-injected mice. At day 26, arginine levels have increased, indicating the accumulation of arginine with the loss of the viral arginase activity. Ornithine levels at day 26 are greatly increased, which is the opposite of uninjected AI-deficient mice that exhibit ornithine deficiency. (d) Relative OAT expression in HDV-mAI- and saline-injected mice. Liver tissue was collected from arg+/− mice at day 14 and day 26, along with arg−/− saline-injected at day 14 of life. OAT expression by RT-qPCR was compared with HDV-mAI-injected mice at day 14 and day 26. Day 14 HDV-mAI-injected mice demonstrate lower levels of OAT expression likely due to near normalization of hepatic arginase activity. Although OAT expression at day 26 is less than two times that of day 26 arg+/− mice, the mechanism of hepatic hyperornithine levels remains unexplained. HDV, helper-dependent adenoviral vector; Het, heterozygous; KO, knockout; mAI, murine AI; OAT, ornithine aminotransferase. Error bars represent mean ± SD.

In addition to hyperammonemia, arginase-deficient mice are characterized by severe hyperargininemia and hypoornithinemia.3 The loss of arginase activity results in the inability of arginine to be metabolized to ornithine and urea, leading to the perturbation of these metabolites. In general, arginine and ornithine levels were examined in the same sets of mice used in some of the plasma ammonia studies (Figure 8b,c). Arg−/− mice injected with HDV-mAI (n = 6) exhibited lower levels of arginine (360.0 ± 219.2 nmol/g protein) at postnatal day 14 compared to saline-injected arg−/− mice (12,064.0 ± 3,868.8 nmol/g protein), (P = 0.006) (n = 6). By day 26, arginine levels had increased up to tenfold in the HDV-mAI injected arg−/− mice (3,688.0 ± 3,711.1 nmol/g protein) (n = 6) indicating an accumulation of arginine coincident with the loss of viral arginase activity (Figure 8b). Saline injected arg−/− mice (n = 6) at day 14 exhibit ornithine deficiency compared to mAI heterozygous mice (n = 6) (13.3 nmol/g protein versus 1,532 ± 50.9 nmol/g protein) (P = 0.0001) (Figure 8c). At day 14, ornithine levels were normalized in the HDV-mAI injected arg−/− mice (n = 6) (1,501.3 ± 354.2 nmol/g protein) (P = 0.915). In contrast, ornithine levels at day 26 in HDV-mAI injected arg−/− mice (n = 6) were greatly increased compared to mAI heterozygous mice (n = 6) (7,202.6 ± 3,880.3 nmol/g protein versus 1,270.0 ± 980.0 nmol/g protein) (P = 0.029) (Figure 8c). This was an unexpected finding, as we anticipated ornithine levels to decrease after the loss of arginase activity, similar to uninjected arginase-deficient mice.

Quantitation of ornithine aminotransferase (OAT) expression was performed to help explain the ornithine levels detected. In addition to arginase, ornithine may be synthesized by OAT from pyrroline-5-carboxylate. Liver tissue from saline-injected arg−/− mice at 14 days of life with tremor phenotype (n = 3), heterozygous mice at 14 days (n = 3) and 26 days of life (n = 3), and HDV-mAI-injected arg−/− mice at 14 days (n = 3) and 26 days of life (n = 3) were compared for expression of OAT activity by reverse transcriptase quantitative real-time PCR and normalized to β-actin expression (Figure 8d). Day 14 of life arg+/− mice had the lowest OAT expression. Although the 14 day arg−/− animals injected with HDV-mAI appear to have normal ornithine metabolism, biochemical data presented here demonstrates that these parameters are not completely normal; OAT expression is increased 3.8 ± 0.4 times that of the day 14 heterozygote. In contrast, the saline-injected day 14 arg−/− is 14.9 ± 0.5 times that of the day 14 heterozygote. Finally, the day 26 mAI arg−/− HDV-injected mouse with measurable biochemical abnormalities and a short remaining life expectancy demonstrates OAT expression 1.9± 0.4 times that of the day 26 heterozygote.

Discussion

Therapies for urea cycle defects and related disorders include (i) substrate reduction or avoidance, (ii) pharmacological treatment with ammonia-conjugating agents, (iii) gene therapy, or (iv) liver transplantation. Although substrate reduction is the mainstay of treatment for urea cycle defects, stem cell replacement and gene therapy are being studied to provide a means of preventing the development of the neurological abnormalities in affected patients. Despite some success with substrate reduction, patients remain at high risk for hyperammonemic crises later in life and the consequences of cumulative neurologic insults.14,15

The liver is an attractive target for gene therapy for several reasons. It is the site of many metabolic pathways and is involved in multiple inborn errors of metabolism. It is a large organ that is readily accessible via the bloodstream through a fenestrated endothelium and in vivo liver transduction with hepatotropic viral vectors is efficient. However, gene therapy has generally been limited by immune responses to either vector-associated antigens, or to the transgene-encoded protein, or by loss of the viral vector by either its disappearance or the cessation of effective transcription. These responses include the development of neutralizing antibodies that block the activity of a secreted protein, or a cytotoxic T lymphocyte response that destroys vector-transduced cells. Decreased transcription associated with extensive methylation of viral promoters is also a major mechanism responsible for a decrease in transgene mRNA levels.16 Whether immune- or promoter-mediated, either response negates the efficacy of gene therapy.

The goal of this study was to determine whether liver-targeted gene therapy could correct the manifestations of arginase deficiency and prolong the life of AI-deficient (knockout) mice that die in the postnatal period. We demonstrated that neonatal gene delivery using a HDV expressing mAI in a tissue-restricted fashion was able to prolong the life of mice with an otherwise lethal phenotype. This correction is accompanied by the production of AI, confirming that successful hepatocyte transduction had occurred. Ammonia levels were decreased at 2 weeks and arginase levels were comparable to arg+/− controls. The murine liver grew over 13-fold its size at birth to 27 days of life. Before death, plasma ammonia was elevated and arginase levels declined to <10% of that of control animals. In essence, the arg−/− mice died from a hyperammonemic crisis due to the loss of AI expression similar to uninjected arg−/− mice, albeit 2 weeks later in life. No immune response was detected to either viral capsid proteins or to AI. Taken together, the data suggest that the loss of HDV-mAI expression is due to the loss or dilution of the DNA from this nonintegrating and nonreplicating viral vector, most likely resulting from the animal's rapid growth and cell division of the mouse liver, the site of viral transduction and transgene expression, and not due to immune-mediated viral or protein loss nor complete silencing of gene expression. Our data on development suggest that the dilution of the viral vector with the massive growth of the liver leaves the organ with inadequate arginase activity to maintain normal or near-normal nitrogen metabolism that was obtainable in the smaller developing mouse causing rising plasma ammonia levels and death.

Death does not appear to be due to an immune-mediated loss of vector expression. We did develop our vector with the strategy of restricting the pattern of expression to the liver to reduce potential host immune responses17,18 and the development of humoral immune responses to the capsid protein or to arginase do not appear to be the cause of loss of AI expression. We had previously demonstrated in other murine models with in utero gene delivery19,20 and others had demonstrated with neonatal gene therapy21,22 that there is a lack of an immune response against either vector-associated or transgene-encoded proteins likely due to the immaturity of immune ontogeny.

Similarly, in the studies reported herein, there was no detectable immune response limiting vector or transgene persistence—again a unique finding when compared to immune-mediated vector loss in adult immunocompetent animals23,24,25,26 or in human clinical trials.27 A decline in viral DNA with a parallel decline in viral RNA with growth of the liver and animal suggests a general dilution and/or loss of viral DNA and, thus, RNA in the background of an expanding cellular, specifically hepatocyte, population. The urea cycle takes place in the periportal hepatocytes, whereas glutamine synthesis is found exclusively in the small perivenous hepatocyte population surrounding the terminal hepatic venule.28,29,30 LacZ staining demonstrates that intravenous HDV administration at day 2 of life results in hepatic transduction throughout the liver lobule with no preference for a specific sublobar location. Death in the face of some but inadequate arginase expression may depend on the location of expression; if residual arginase activity was located outside of the periportal hepatocytes (i.e., perivenous zone) in the liver acinus, then this might not reflect the true effective urea cycle residual arginase activity.

Metabolite levels including serum arginine, ornithine, and ammonia were assessed as endpoints. In vector-administered arg−/− mice, we saw ammonia levels decline to near normal values and also a decline in arginine. These levels increased with the loss of arginase activity, mimicking the onset of symptoms in arginase deficiency. When correlated with arginase activity data, these results suggest that having as low as 20% endogenous arginase activity is enough to maintain normal ammonia levels. However, if activity drops <10% of normal, the urea cycle does not function well, and ammonia levels begin to increase. Understanding these minimal functional levels will be of great importance as we begin to strategize approaches for gene therapy in human AI-deficient patients.

Although ornithine levels were normalized in vector-administered arg−/− mice, instead of decreasing upon loss of arginase expression, levels continued to rise up to sevenfold higher than normal levels. In wild-type mice, arginase in conjunction with OAT can produce ornithine. OAT reversibly catalyzes the conversion of ornithine and α-ketoglutarate to glutamic-γ-semialdehyde and glutamate. During the first 2 weeks of life in mice, OAT in the gastrointestinal tract is the main source of ornithine, at which point OAT levels drop and arginase is responsible for ornithine production.31 It is possible that the expression of vector-administered AI specifically in the liver of arg−/− mice perturbs the regulation of OAT, either by decreasing OAT or pushing the equilibrium of the reaction toward production of ornithine. Studies conducted herein demonstrate that OAT expression is increased in arg−/− mice and that HDV-mAI-treated animals have lower levels of OAT at day 14. OAT expression is increased as animals approach day 27, likely due to loss of vector and thus arginase expression. While our data demonstrate a trend toward normalization of OAT in young vector-treated animals, the modestly elevated OAT expression does not explain the substantially elevated ornithine levels in HDV-mAI-injected arg−/− mice at day 26. Thus, our hypothesis that diminished liver ornithine is responsible for the development of hyperammonemia, must be called into question, or at least be explained by a compartmentalization phenomenon in which the ornithine produced is not in the same periportal area of the liver in which the urea cycle occurs.

Hyperornithinemia in humans is caused by OAT deficiency and results in gyrate atrophy of the choroid and retina, presenting as night blindness.32 OAT deficient mice, which survive into adulthood, have ornithine levels 10- to 15-fold higher than normal and exhibit slowly progressive retinal degeneration.33 These data suggest that the sevenfold increase in ornithine levels we detected are not contributing to the death of the HDV-mAI injected arg−/− mice.

Taken together, these results suggest that early postnatal treatment of urea cycle defects is possible but may require a vector where the transgene genome is not lost with cellular division. This loss, which at least in part is due to the episomal location of viral DNA, may be prevented with a hybrid adenoviral vector, that is one based on adenovirus to provide efficient gene transfer and an integrating vector to provide stable gene expression and prevent loss of the transgene.34,35,36 Using integrative vectors, the transduced cellular genome ensures permanent expression, regardless of the number of subsequent cellular divisions. This is likely a major requirement if gene therapy is to be applicable to metabolic diseases that present early in childhood or in the neonatal period. However, a major limitation of integrating vectors is the risk of insertional mutagenesis, illustrated by the development of a leukemia-like illness in some subjects of a gene therapy trial for severe X-linked SCID.37 Further murine studies using recombinant adeno-associated viral vectors are currently under way in which we have detected reporter gene expression for up to 27 months (G.S. Lipshutz, unpublished results). Alternatively, mice could be administered a second dose of HDV-mAI to transduce additional hepatocytes as they develop and grow in size. This form of “sequential gene therapy,” which we are currently examining, may result in significant abrogative immune responses not only to viral capsid proteins but also to AI. Serotype switching may be one strategy to overcome existing adenoviral neutralizing antibodies.38,39

In conclusion, we believe that this study demonstrates that in vivo gene delivery to the liver using viral vectors in neonatal mice with AI deficiency is a valid therapeutic strategy that deserves further investigation and with further development may be considered for future clinical application. Addressing the effect of cellular division will be important in any strategy treating neonates or directed to organs with significant cellular turnover.

Materials and Methods

Generation of HDVs. mAI cDNA was obtained by PCR amplification. Total RNA was isolated and cDNA was generated by using oligo dT primers and reverse transcriptase. Arginase-specific primers (forward: 5′-ATG AGC TCC AAG CCA AAG TCC TTA GAG A-3′ and reverse: 5′-TCA CTT AGG TGG TTT AAG GTA GTC A-3′) were used. Amplification was performed for 30 cycles (94 for 30 seconds, 60 for 30 seconds, 72 for 1 minute) after denaturing for 2 minutes at 94. Extension was continued at 72 for 10 minutes at completion. mAI cDNA was cloned into pCRII-TOPO (Invitrogen, Carlsbad, CA) and sequenced using vector-specific primers and compared to GENBANK. Subsequently, mAI cDNA with Not I linkers was cloned into pPEPCK vector downstream of the phosphoenolpyruvate carboxykinase (PEPCK) promoter and upstream of the WPRE element to generate pPEPCK-mAI. This plasmid was then enzymatically digested with AscI, and the insert containing the PEPCK promoter, mAI ORF, and the WPRE was subcloned into pD28E4 to produce pD28E4-mAI (Figure 1). The LacZ cDNA was cloned into the Not I site of the pPEPCK plasmid. The PEPCK-LacZ expression cassette was digested with Asc I and inserted into the pD21.7E4 plasmid.40

The HDV-PEPCK-mAI-WPRE and the HDV-PEPCK-LacZ-WPRE were produced using the helper virus AdNG163R-241 and 116 producer cells, as described in detail elsewhere.41,42 The HDV vectors were purified by triple CsCl ultracentrifugation and formulated in 10 mmol/l Tris-HCl, pH 8.0 supplemented with glycerol to 10%. Virion DNA was extracted for analysis by Southern blot hybridization and the levels of helper virus contamination determined by phosphoimager analysis were found to be <0.05% for both vector preparations. Physical vector titer was determined spectrophotometrically and expressed as vector particles (vp)/ml, as described in detail elsewhere.43

Mouse injection and sample collection. All handling of animals was performed in accordance with the University of California, Los Angeles (UCLA) Chancellor's Committee for Animal Research guidelines. AI knockout mice were generated within our laboratory and have been previously described.3 Heterozygous mAI mice were mated to produce litters containing wild type, arg+/−, and arg−/− mice. All pups of a litter were injected intravenously via the superficial temporal vein between 2 and 3 days of life with either pharmaceutical grade saline or 5 × 109 viral particles of HDV-mAI per gram body weight. Mice were genotyped on postnatal day 7 using PCR.3 A three-primer PCR was performed (mAI forward: 5′-AAC CAG CAC CTC TAA GGT CTA TGG-3′; Neo forward: 5′-GCC CAT TCG ACC ACC AAG-3′; Exon 5 Reverse: 5′-ACG ATG TCT TTG GCA GAT ATG C-3′) using Pfu turbo (Stratagene, La Jolla, CA) and Platinum Taq (Invitrogen) for 35 cycles (94 for 30 seconds, 60 for 30 seconds, and 72 for 3 minutes).

Wild-type and heterozygous mice were euthanized at predetermined time points to follow localization and expression of HDV-mAI. A total of 15 arg−/− mice injected with HDV-mAI were observed until they showed a decline in health. In addition, some were removed and euthanized at predetermined time points to compare with saline-injected mice. Multiple tissues were removed for use in examining HDV-mAI expression and amino acid analysis studies.

HDV-mAI expression. mRNA expression was quantified by isolating RNA from tissue (RNeasy Mini Kit; Qiagen, Valencia, CA). cDNA was then synthesized using the First Strand cDNA Synthesis Kit (Fermentas, Hanover, MD), and reverse transcriptase PCR was performed using primers located in the 3′ end of mAI gene (mAI int forward: 5′-GCT TGC GAG ACG TAG ACC-3′) and the 5′ region of the WPRE (WPRE rev: 5′-GTT GCG TCA GCA AAC ACA GTG-3′) for 35 cycles (94 for 30 seconds, 57 for 30 seconds and 72 for 1 minute). β-actin PCR was performed as a control using (β-actin forward: 5′-GTG GGG CGC CCC AGG CAC CA-3′; β-actin reverse: 5′-CTC CTT AAT GTC ACG CAC GAT TTC-3′) for 35 cycles (94 for 30 seconds, 60 for 30 seconds and 72 for 90 seconds).44

Western blot. Protein analysis of HDV-mAI expression was performed first by homogenizing 40 mg of liver in 1 ml of cold RIPA buffer containing 1× HALT protease inhibitor cocktail (Pierce Biotechnology, Rockford, IL). The total protein content was determined using a Lowry-based assay (Bio-Rad, Hercules, CA) with the absorbance measured at 750 nm. Proteins were denatured with loading dye that consisted of 95:5 Laemmli Buffer:β-mercaptoethanol (Bio-Rad) and 25 µg of each sample was subsequently loaded on a precast 12% PAGE gel for separation, which was performed with a MiniProtean II Cell at 115V for 2 hours. PVDF membranes were washed for 1 minute in methanol and equilibrated in transfer buffer (Bio-Rad) for 30 minutes. Transfer to the PVDF was performed overnight at 25V in 4 °C. The membrane was blocked in 5% dry milk/PBST solution for at least 10 hours at 4 °C and incubated for 2 hours at room temperature with CJ rabbit-anti-rat AI antibody45 followed by 1 hour room temperature incubation with goat-anti-rabbit-HRP conjugated secondary antibody (Santa Cruz Biotechnology, Santa Cruz, CA).27 To detect the proteins, the membrane was incubated in a chemiluminescent substrate (Pierce Biotechnology) and exposed to film.

As a loading control, protein analysis of β-actin expression was performed using the same samples as previously mentioned. Accordingly, the membrane was stripped by incubation in 30 ml of Restore Western Blot Stripping Buffer (Pierce Biotechnology) for 15 minutes at room temperature and then washed. Subsequently, the membrane was incubated with mouse monoclonal β-actin antibody (Abcam, Cambridge, MA) followed by antimouse-HRP-conjugated secondary antibody (BD Pharmingen, Franklin, NJ).

Virus biodistribution. Transduction efficiencies of neonatal mouse liver after vector administration of the LacZ reporter gene were determined by real-time PCR.46 A two-primer real-time PCR was performed (LacZ forward: 5′-CCC AAC TTA ATC GCC TTG CA-3′; LacZ reverse: 5′-GCG GGC CTC TTC GCT ATT-3′) using Platinum SYBR Green (Invitrogen) for 40 cycles (95 °C for 15 seconds, 60 °C for 1 minute) after denaturing for 10 minutes at 95 °C. A standard curve ranging from 102 to 107 copies of LacZ per cell was generated by serial dilution of plasmid pEF1/Myc-His/LacZ (9.2 kb) (Invitrogen) in TE and genomic DNA. As an internal control, a β-actin qPCR was also performed. A two-primer real-time PCR (β-actin forward: 5′-GTG GGC CGC TCT AGG CAC CA-3′; β-actin reverse: 5′-CGG TTG GCC TTA GGG TTC AGG GGG-3′) using Platinum SYBR Green (Invitrogen) was performed for 40 cycles (95 °C for 15 seconds, 59 °C for 10 seconds, and 72 °C for 15 seconds) after denaturing for 10 minutes at 95 °C. Relative expression ratios of β-actin were used to correct for sample variability.

Viral expression in liver. X-gal staining was performed on 5-µm tissue sections to determine location of hepatic expression. Tissues were fixed and slides were placed in X-gal solution overnight at 37 °C, as previously described.19 Tissues were counterstained with hematoxylin (Sigma Chemical, St Louis, MO).

Viral mRNA expression was determined by RT-qPCR. After isolating RNA and synthesizing cDNA as previously discussed, real-time PCR was performed using the same LacZ primers and conditions described. For the qPCR, 2 µl of the RT-product was utilized.

Determination of arginase levels. Arginase activity of lysates was measured by a colorimetric assay described previously.30 Briefly, the livers were homogenized in 40 µl of 0.1% Triton X-100 and HALT protease inhibitor cocktail per mg tissue. 10 µl of lysate was diluted with water to a final volume of 100 µl, and added to 100 µl of 25 mmol/l Tris-HCl (pH 7.5) and 20 µl of 10 mmol/l MnCl2. Arginase was activated by heating the sample at 56 °C for 10 minutes. 100 µl of 0.5 mol/l L-arginine (pH 7.9) was added and incubated at 37 °C for 1 hour. The conversion of arginine to urea was stopped by the addition of 900 µl of acid mix [1:3:7 mixture of H2SO4 (96%):H3PO4 (85%):water]. 40 µl of 9% 1-phenyl-1,2-propanedione-2-oxime (ISPF) dissolved in ethanol was added and heated to 95 °C for 30 minutes for color development. Each sample was diluted 1:6 with water and 200 µl measured for absorbance at 540 nm. Urea standards ranging from 0.03 to 0.5 µmol were used to create a calibration curve for each assay. To normalize the arginase activity measured, as previously mentioned, the total protein content was then assayed using the Lowry-based assay (Bio-Rad), as described.

Examination of immune responses to adenovirus and arginase. Immune response to viral capsid protein was measured by antiadenovirus ELISA. Ninety-six well immunoplates (NUNC, Rochester, NY) were coated with 2 × 108 viral particles of HDV-mAI that were prepared in 75 µl of 50 mmol/l carbonate buffer (pH 9.6). To inactivate the virus, the plates were exposed to UV light for 30 minutes. Following an overnight incubation at 4 °C and washing with 1 × PBS, 200 µl of blocking reagent (PBS with 5% FCS) was added to each well and incubated at 37 °C for 2 hours. Meanwhile, twofold dilutions of plasma samples from experimental (neonatally injected with HDV-LacZ now at 24 days of life) and control groups, beginning from 1:40, were prepared in 1% BSA/PBST. After the plates were washed, 100 µl of diluted plasma was added in duplicate and subsequently incubated for 2 hours at 37 °C. With a dilution of 1:1,000 of HRP-conjugated-goat-antimouse secondary antibody (BD Pharmingen), 50 µl was added and incubated for 1 hour at 37 °C. The plates were washed, as mentioned previously, and incubated with 50 µl of OPD substrate for up to 10 minutes. Color development was stopped by adding 50 µl of 2.5 mol/l sulfuric acid and was measured for absorbance at 492 nm; background was defined as the absorbance measured in the absence of primary and secondary antibodies. Antiarginase ELISA was similarly performed, but the immunoplates were instead coated with 50 ng of human AI (Alexis Biochemicals, San Diego, CA). For these studies, serum was collected from day 26 arg−/− mice, which received HDV-mAI at 2 days of life. A positive control antiadenovirus antibody was generated by intraperitoneally administering a naive 8-week-old adult mouse with 5 × 1010 HDV-LacZ viral particles mixed with alum (Pierce Biotechnology). Blood was collected 4 weeks later and serum was removed and stored at –80 °C until needed. A mouse antiarginase antibody (BD Pharmingen) was used as positive control for the antiarginase ELISA. Results were presented as average ±SD.

Examination of cellular responses was determined by Hematoxylin and Eosin staining of liver sections at days of life 8, 12, 18, 24, 28, 32, 40 for both saline-injected and HDV-injected mice (see Supplementary Materials and Methods).

Plasma ammonia analysis. Plasma of arg−/− or arg+/− mice injected with saline or HDV-mAI was collected at postnatal day 14 or 26 (HDV-mAI only), as described previously.31 Briefly, blood was drawn by cardiac puncture or by retro-orbital bleeding and collected in nonheparinized microhematocrit capillary tubes (Fisher Scientific, Pittsburgh, PA). Plasma was separated from red blood cells by centrifugation, and assayed immediately with a plasma ammonia assay kit (Sigma Chemical) according to the manufacturer's instructions. For each sample, 25 µl was added to water for a final volume of 100 µl for a 1:4 dilution. The absorbance was measured spectrophotometrically at 340 nm and ammonia concentrations were calculated according to the noncompetitive reaction equation.

Amino acid analysis. Liver arginine and ornithine levels were measured in arg−/−or arg+/− mice injected with saline or HDV-mAI and collected at postnatal day 14 or 26 (HDV-mAI only). For each sample, two flash-frozen tissues were combined, weighed, and homogenized in water to a concentration of 0.2 g/ml. The supernatant was then removed and sonicated, 1 ml was removed and 70 mg of sulfosalicylic acid was added. The samples were then centrifuged. Supernatants were removed and sent on dry ice to the laboratory of Stephen Goodman (University of Colorado Health Sciences Center, Denver, CO) for amino acid analysis on a Beckman System 6300 Amino Acid Analyzer utilizing a 3-Lithium buffer method, as described previously.31 Results were reported in nanomoles per gram of tissue.

Determination of ornithine amino transferase activity. Relative OAT expression in the livers of vector-treated and untreated knockout mice was determined by RT-qPCR. RNA was isolated from livers (RNeasy Mini Kit; Qiagen), and cDNA was synthesized (First Strand cDNA Synthesis Kit; Fermentas, Hanover, MD). Real-time PCR was then performed using platinum SYBR Green (Invitrogen) and murine OAT specific primers (forward 5′-GGG CTC TTG TGA AAC TCT GC-3′ and reverse 5′-AGA TGG GTC CGT TTC TCC TT-3′) for 40 cycles (95 °C for 15 seconds, 56 °C for 30 seconds, 72 °C for 15 seconds), after denaturing for 10 minutes at 95 °C.47 β-Actin PCR was used to standardize the relative mOAT expression ratios.48

Statistical analysis. Mouse survival was compared by log rank. One-way ANOVA was calculated for statistical significance for arginase activity, ammonia, and amino acid levels. Statistical calculations were performed using the SPSS version 17.0 statistical software package (SPSS, Chicago, IL). Data are presented as mean ± SD unless stated otherwise.

SUPPLEMENTARY MATERIALFigure S1. Histology of liver of mice injected with HDV. Liver sections (400x) of HDV-injected (A) or saline-injected (B) mice were examined for cellular infiltrates. Mice were examined on day of life 8 (panel 1), 12 (panel 2), 18 (panel 3), 24 (panel 4), 27 (panel 5), 30 (panel 6), and 40 (panel 7). All mice were injected intravenously on the second day of life. No infiltrates were detected in either saline or HDV-injected mice (n≥3 per group).Materials and Methods.

Supplementary Material

Figure S1.

Histology of liver of mice injected with HDV. Liver sections (400x) of HDV-injected (A) or saline-injected (B) mice were examined for cellular infiltrates. Mice were examined on day of life 8 (panel 1), 12 (panel 2), 18 (panel 3), 24 (panel 4), 27 (panel 5), 30 (panel 6), and 40 (panel 7). All mice were injected intravenously on the second day of life. No infiltrates were detected in either saline or HDV-injected mice (n≥3 per group).

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Materials and Methods.
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Acknowledgments

This work was supported by NIH grants HD057555-02A1 to G.S.L., HD-06576 HD-04612 to S.D.C., and to K99HL088692-01 to N.B.-P. Additional support to G.S.L. and W.W.G. was provided by a Stein–Oppenheimer award. C.L.G. was supported by the T32 GM008243 UCLA Intercampus Medical Genetics NIH training grant. Additional support was provided by the UCLA Mental Retardation Research Center. We thank Tamara Horwich for her assistance with the statistical analysis.

REFERENCES

  1. Iyer R, Jenkinson CP, Vockley JG, Kern RM, Grody WW., and , Cederbaum S. The human arginases and arginase deficiency. J Inherit Metab Dis. 1998;21 Suppl. 1:86–100. doi: 10.1023/a:1005313809037. [DOI] [PubMed] [Google Scholar]
  2. Vockley JG, Jenkinson CP, Shukla H, Kern RM, Grody WW., and , Cederbaum SD. Cloning and characterization of the human type II arginase gene. Genomics. 1996;38:118–123. doi: 10.1006/geno.1996.0606. [DOI] [PubMed] [Google Scholar]
  3. Iyer RK, Yoo PK, Kern RM, Rozengurt N, Tsoa R, O'Brien WE, et al. Mouse model for human arginase deficiency. Mol Cell Biol. 2002;22:4491–4498. doi: 10.1128/MCB.22.13.4491-4498.2002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Benihoud K, Yeh P., and , Perricaudet M. Adenovirus vectors for gene delivery. Curr Opin Biotechnol. 1999;10:440–447. doi: 10.1016/s0958-1669(99)00007-5. [DOI] [PubMed] [Google Scholar]
  5. Brunetti-Pierri N, Stapleton GE, Law M, Breinholt J, Palmer DJ, Zuo Y, et al. Efficient, long-term hepatic gene transfer using clinically relevant HDAd doses by balloon occlusion catheter delivery in nonhuman primates. Mol Ther. 2009;17:327–333. doi: 10.1038/mt.2008.257. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Morral N, Parks RJ, Zhou H, Langston C, Schiedner G, Quinones J, et al. High doses of a helper-dependent adenoviral vector yield supraphysiological levels of alpha1-antitrypsin with negligible toxicity. Hum Gene Ther. 1998;9:2709–2716. doi: 10.1089/hum.1998.9.18-2709. [DOI] [PubMed] [Google Scholar]
  7. Brunetti-Pierri N, Nichols TC, McCorquodale S, Merricks E, Palmer DJ, Beaudet AL, et al. Sustained phenotypic correction of canine hemophilia B after systemic administration of helper-dependent adenoviral vector. Hum Gene Ther. 2005;16:811–820. doi: 10.1089/hum.2005.16.811. [DOI] [PubMed] [Google Scholar]
  8. Brunetti-Pierri N, Stapleton GE, Palmer DJ, Zuo Y, Mane VP, Finegold MJ, et al. Pseudo-hydrodynamic delivery of helper-dependent adenoviral vectors into non-human primates for liver-directed gene therapy. Mol Ther. 2007;15:732–740. doi: 10.1038/sj.mt.6300102. [DOI] [PubMed] [Google Scholar]
  9. Brunetti-Pierri N, Ng T, Iannitti DA, Palmer DJ, Beaudet AL, Finegold MJ, et al. Improved hepatic transduction, reduced systemic vector dissemination, and long-term transgene expression by delivering helper-dependent adenoviral vectors into the surgically isolated liver of nonhuman primates. Hum Gene Ther. 2006;17:391–404. doi: 10.1089/hum.2006.17.391. [DOI] [PubMed] [Google Scholar]
  10. Fausto N., and , Campbell JS. The role of hepatocytes and oval cells in liver regeneration and repopulation. Mech Dev. 2003;120:117–130. doi: 10.1016/s0925-4773(02)00338-6. [DOI] [PubMed] [Google Scholar]
  11. Magami Y, Azuma T, Inokuchi H, Kokuno S, Moriyasu F, Kawai K, et al. Cell proliferation and renewal of normal hepatocytes and bile duct cells in adult mouse liver. Liver. 2002;22:419–425. doi: 10.1034/j.1600-0676.2002.01702.x. [DOI] [PubMed] [Google Scholar]
  12. Wang Z, Zhu T, Qiao C, Zhou L, Wang B, Zhang J, et al. Adeno-associated virus serotype 8 efficiently delivers genes to muscle and heart. Nat Biotechnol. 2005;23:321–328. doi: 10.1038/nbt1073. [DOI] [PubMed] [Google Scholar]
  13. Cunningham SC, Dane AP, Spinoulas A, Logan GJ., and , Alexander IE. Gene delivery to the juvenile mouse liver using AAV2/8 vectors. Mol Ther. 2008;16:1081–1088. doi: 10.1038/mt.2008.72. [DOI] [PubMed] [Google Scholar]
  14. Bachmann C. Outcome and survival of 88 patients with urea cycle disorders: a retrospective evaluation. Eur J Pediatr. 2003;162:410–416. doi: 10.1007/s00431-003-1188-9. [DOI] [PubMed] [Google Scholar]
  15. Enns GM, Berry SA, Berry GT, Rhead WJ, Brusilow SW., and , Hamosh A. Survival after treatment with phenylacetate and benzoate for urea-cycle disorders. N Engl J Med. 2007;356:2282–2292. doi: 10.1056/NEJMoa066596. [DOI] [PubMed] [Google Scholar]
  16. Brooks AR, Harkins RN, Wang P, Qian HS, Liu P., and , Rubanyi GM. Transcriptional silencing is associated with extensive methylation of the CMV promoter following adenoviral gene delivery to muscle. J Gene Med. 2004;6:395–404. doi: 10.1002/jgm.516. [DOI] [PubMed] [Google Scholar]
  17. Ding E, Hu H, Hodges BL, Serra D, Xu F, Chen YT, et al. Efficacy of gene therapy for a prototypical lysosomal storage disease (GSD-II) is critically dependent on vector dose, transgene promoter, and the tissues targeted for vector transduction. Mol Ther. 2002;5:436–446. doi: 10.1006/mthe.2002.0563. [DOI] [PubMed] [Google Scholar]
  18. Pastore L, Morral N, Zhou H, Garcia R, Parks RJ, Kochanek S, et al. Use of a liver-specific promoter reduces immune response to the transgene in adenoviral vectors. Hum Gene Ther. 1999;10:1773–1781. doi: 10.1089/10430349950017455. [DOI] [PubMed] [Google Scholar]
  19. Lipshutz GS, Flebbe-Rehwaldt L., and , Gaensler KM. Reexpression following readministration of an adenoviral vector in adult mice after initial in utero adenoviral administration. Mol Ther. 2000;2:374–380. doi: 10.1006/mthe.2000.0136. [DOI] [PubMed] [Google Scholar]
  20. Lipshutz GS, Gruber CA, Cao Y, Hardy J, Contag CH., and , Gaensler KM. In utero delivery of adeno-associated viral vectors: intraperitoneal gene transfer produces long-term expression. Mol Ther. 2001;3:284–292. doi: 10.1006/mthe.2001.0267. [DOI] [PubMed] [Google Scholar]
  21. Xu L, Mei M, Haskins ME, Nichols TC, O'donnell P, Cullen K, et al. Immune response after neonatal transfer of a human factor IX-expressing retroviral vector in dogs, cats, and mice. Thromb Res. 2007;120:269–280. doi: 10.1016/j.thromres.2006.09.010. [DOI] [PubMed] [Google Scholar]
  22. Traas AM, Wang P, Ma X, Tittiger M, Schaller L, O'donnell P, et al. Correction of clinical manifestations of canine mucopolysaccharidosis I with neonatal retroviral vector gene therapy. Mol Ther. 2007;15:1423–1431. doi: 10.1038/sj.mt.6300201. [DOI] [PubMed] [Google Scholar]
  23. Jooss K., and , Chirmule N. Immunity to adenovirus and adeno-associated viral vectors: implications for gene therapy. Gene Ther. 2003;10:955–963. doi: 10.1038/sj.gt.3302037. [DOI] [PubMed] [Google Scholar]
  24. Muruve DA. The innate immune response to adenovirus vectors. Hum Gene Ther. 2004;15:1157–1166. doi: 10.1089/hum.2004.15.1157. [DOI] [PubMed] [Google Scholar]
  25. Muruve DA, Cotter MJ, Zaiss AK, White LR, Liu Q, Chan T, et al. Helper-dependent adenovirus vectors elicit intact innate but attenuated adaptive host immune responses in vivo. J Virol. 2004;78:5966–5972. doi: 10.1128/JVI.78.11.5966-5972.2004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Zaiss AK., and , Muruve DA. Immune responses to adeno-associated virus vectors. Curr Gene Ther. 2005;5:323–331. doi: 10.2174/1566523054065039. [DOI] [PubMed] [Google Scholar]
  27. Manno CS, Pierce GF, Arruda VR, Glader B, Ragni M, Rasko JJ, et al. Successful transduction of liver in hemophilia by AAV-Factor IX and limitations imposed by the host immune response. Nat Med. 2006;12:342–347. doi: 10.1038/nm1358. [DOI] [PubMed] [Google Scholar]
  28. Haussinger D., and , Gerok W. Hepatocyte heterogeneity in glutamate uptake by isolated perfused rat liver. Eur J Biochem. 1983;136:421–425. doi: 10.1111/j.1432-1033.1983.tb07759.x. [DOI] [PubMed] [Google Scholar]
  29. Haussinger D, Sies H., and , Gerok W. Functional hepatocyte heterogeneity in ammonia metabolism. The intercellular glutamine cycle. J Hepatol. 1985;1:3–14. doi: 10.1016/s0168-8278(85)80063-5. [DOI] [PubMed] [Google Scholar]
  30. Gebhardt R., and , Mecke D. Heterogeneous distribution of glutamine synthetase among rat liver parenchymal cells in situ and in primary culture. Embo J. 1983;2:567–570. doi: 10.1002/j.1460-2075.1983.tb01464.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Wang T, Lawler AM, Steel G, Sipila I, Milam AH., and , Valle D. Mice lacking ornithine-delta-aminotransferase have paradoxical neonatal hypoornithinaemia and retinal degeneration. Nat Genet. 1995;11:185–190. doi: 10.1038/ng1095-185. [DOI] [PubMed] [Google Scholar]
  32. Takki K., and , Simell O. Genetic aspects in gyrate atrophy of the choroid and retina with hyperornithinaemia. Br J Ophthalmol. 1974;58:907–916. doi: 10.1136/bjo.58.11.907. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Wang T, Milam AH, Steel G., and , Valle D. A mouse model of gyrate atrophy of the choroid and retina. Early retinal pigment epithelium damage and progressive retinal degeneration. J Clin Invest. 1996;97:2753–2762. doi: 10.1172/JCI118730. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Ehrhardt A, Yant SR, Giering JC, Xu H, Engler JA., and , Kay MA. Somatic integration from an adenoviral hybrid vector into a hot spot in mouse liver results in persistent transgene expression levels in vivo. Mol Ther. 2007;15:146–156. doi: 10.1038/sj.mt.6300011. [DOI] [PubMed] [Google Scholar]
  35. Picard-Maureau M, Kreppel F, Lindemann D, Juretzek T, Herchenröder O, Rethwilm A, et al. Foamy virus—adenovirus hybrid vectors. Gene Ther. 2004;11:722–728. doi: 10.1038/sj.gt.3302216. [DOI] [PubMed] [Google Scholar]
  36. Murphy SJ, Chong H, Bell S, Diaz RM., and , Vile RG. Novel integrating adenoviral/retroviral hybrid vector for gene therapy. Hum Gene Ther. 2002;13:745–760. doi: 10.1089/104303402317322302. [DOI] [PubMed] [Google Scholar]
  37. Hacein-Bey-Abina S, Von Kalle C, Schmidt M, McCormack MP, Wulffraat N, Leboulch P, et al. LMO2-associated clonal T cell proliferation in two patients after gene therapy for SCID-X1. Science. 2003;302:415–419. doi: 10.1126/science.1088547. [DOI] [PubMed] [Google Scholar]
  38. Mack CA, Song WR, Carpenter H, Wickham TJ, Kovesdi I, Harvey BG, et al. Circumvention of anti-adenovirus neutralizing immunity by administration of an adenoviral vector of an alternate serotype. Hum Gene Ther. 1997;8:99–109. doi: 10.1089/hum.1997.8.1-99. [DOI] [PubMed] [Google Scholar]
  39. Mastrangeli A, Harvey BG, Yao J, Wolff G, Kovesdi I, Crystal RG, et al. “Sero-switch” adenovirus-mediated in vivo gene transfer: circumvention of anti-adenovirus humoral immune defenses against repeat adenovirus vector administration by changing the adenovirus serotype. Hum Gene Ther. 1996;7:79–87. doi: 10.1089/hum.1996.7.1-79. [DOI] [PubMed] [Google Scholar]
  40. Brunetti-Pierri N, Palmer DJ, Mane V, Finegold M, Beaudet AL., and , Ng P. Increased hepatic transduction with reduced systemic dissemination and proinflammatory cytokines following hydrodynamic injection of helper-dependent adenoviral vectors. Mol Ther. 2005;12:99–106. doi: 10.1016/j.ymthe.2005.03.001. [DOI] [PubMed] [Google Scholar]
  41. Palmer D., and , Ng P. Improved system for helper-dependent adenoviral vector production. Mol Ther. 2003;8:846–852. doi: 10.1016/j.ymthe.2003.08.014. [DOI] [PubMed] [Google Scholar]
  42. Palmer DJ., and , Ng P. Physical and infectious titers of helper-dependent adenoviral vectors: a method of direct comparison to the adenovirus reference material. Mol Ther. 2004;10:792–798. doi: 10.1016/j.ymthe.2004.06.1013. [DOI] [PubMed] [Google Scholar]
  43. Ng P, Parks RJ., and , Graham FL. Preparation of helper-dependent adenoviral vectors. Methods Mol Med. 2002;69:371–388. doi: 10.1385/1-59259-141-8:371. [DOI] [PubMed] [Google Scholar]
  44. Tanikawa M, Harada T, Mitsunari M, Onohara Y, Iwabe T., and , Terakawa N. Expression of c-kit messenger ribonucleic acid in human oocyte and presence of soluble c-kit in follicular fluid. J Clin Endocrinol Metab. 1998;83:1239–1242. doi: 10.1210/jcem.83.4.4746. [DOI] [PubMed] [Google Scholar]
  45. Spector EB, Kern RM, Haggerty DF., and , Cederbaum SD. Differential expression of multiple forms of arginase in cultured cells. Mol Cell Biochem. 1985;66:45–53. doi: 10.1007/BF00231822. [DOI] [PubMed] [Google Scholar]
  46. Koponen JK, Turunen AM., and , Yla-Herttuala S. Escherichia coli DNA contamination in AmpliTaq Gold polymerase interferes with TaqMan analysis of lacZ. Mol Ther. 2002;5:220–222. doi: 10.1006/mthe.2002.0548. [DOI] [PubMed] [Google Scholar]
  47. Ventura G, De Bandt JP, Segaud F, Perret C, Robic D, Levillain O, et al. Overexpression of ornithine aminotransferase: consequences on amino acid homeostasis. Br J Nutr. 2008;101:843–851. doi: 10.1017/S0007114508043389. [DOI] [PubMed] [Google Scholar]
  48. Deignan JL, Livesay JC, Shantz LM, Pegg AE, O'Brien WE, Iyer RK, et al. Polyamine homeostasis in arginase knockout mice. Am J Physiol Cell Physiol. 2007;293:C1296–C1301. doi: 10.1152/ajpcell.00393.2006. [DOI] [PubMed] [Google Scholar]

Associated Data

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Supplementary Materials

Figure S1.

Histology of liver of mice injected with HDV. Liver sections (400x) of HDV-injected (A) or saline-injected (B) mice were examined for cellular infiltrates. Mice were examined on day of life 8 (panel 1), 12 (panel 2), 18 (panel 3), 24 (panel 4), 27 (panel 5), 30 (panel 6), and 40 (panel 7). All mice were injected intravenously on the second day of life. No infiltrates were detected in either saline or HDV-injected mice (n≥3 per group).

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Materials and Methods.
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