Introduction

Lysosomal storage diseases (LSD) are a varied group of inherited metabolic disorders that share a common pathophysiology in which very low or absent activity of a specific enzyme causes gradual lysosomal accumulation of uncatabolized macromolecular substrate, including cerebroside, ganglioside, glycosphingolipid, glycogen, glycoprotein, or mucopolysaccharide¹. LSDs have a worldwide incidence of approximately 1 in 8000 live births. Of these, approximately 1/3 are one of the mucopolysaccharidoses (MPS), a deficiency in glycosaminoglycan (GAG) catabolism. MPS I is caused by a loss of activity of the enzyme -L-iduronidase, the lysosomal protein required to initiate the breakdown of sulfated GAGs dermatan sulfate and heparan sulfate². More than 75 mutations have been identified in the human IDUA gene³. Homozygosity for some mutations (e.g., W402X and Q70X) results in a total loss of enzyme activity and the most severe phenotype, Hurler syndrome. By about 2 years of age, severely affected children begin to exhibit characteristic features of the disease, including growth delay, coarsening facial features, excess urinary GAG, hepatosplenomegaly, skeletal abnormalities, and neurological deficits. Affected children typically die by age 10 years due to cardiac failure or fatal recurrent respiratory infection².

Proposed therapies for MPS I have been based on early cellular biochemical analyses⁴ and include enzyme replacement^5,6, infusion of normal leukocytes⁷, and allogeneic bone marrow transplantation^8,9,10,11. It has been shown that a small portion of newly synthesized enzyme that normally traffics to the lysosomal compartment escapes into the extracellular environment, where it can be endocytosed by means of association with the mannose-6-phosphate receptor⁴. Reconstitution of IDUA activity as evidenced by increased peripheral enzyme levels, improved longevity, and sustained cognitive development has resulted from allogeneic bone marrow transplant, thus indicating that introduction of normal gene sequences will benefit the patient^9,10,11. Difficulties with availability of matched donors and inherent allogeneic transplant complications continue to motivate the search for alternative therapies^10,11.

The success of allogeneic marrow transplant and enzyme therapy has fueled anticipation that some LSDs may be amenable to treatment by gene therapy^12,13. To address this possibility, mouse and dog models of MPS VII have allowed tremendous insight into the potential for treatment of LSD by gene transfer^14,15,16. Importantly, it has been possible to express high levels of -glucuronidase (GUSB) activity, with concomitant clearance of storage material and normalization of functional deficits after intravenous AAV-GUSB injection in 1-day-old MPS VII mice^17,18. Retroviral vector-mediated GUSB gene transfer in MPS VII dogs has been shown to diminish systemic effects of the disease and led to a striking recovery of musculoskeletal deficiencies in these animals¹⁹. Recently, injection of a recombinant murine retrovirus carrying the canine GUSB sequence into MPS VII mice led to high levels of gene transfer and expression that were sufficient to eliminate lysosomal storage pathology in a wide array of peripheral tissues and numerous subregions of the brain¹⁶.

A primary goal of gene therapy for LSD is preservation of neurological function. MPS VII mice treated with multiple systemic infusions of GUSB enzyme demonstrated learning and memory trends in the Morris water maze test that were identical to those of normal controls²⁰. Subsequently, it has been shown that a direct injection of AAV-GUSB into the brain of MPS VII mice could result in stable enzyme expression in this tissue, which was sufficient to mediate significant improvement in most parameters of the Morris water maze test²¹. Direct injection of a feline immunodeficiency virus vector carrying GUSB sequence into the striatum of 4-month-old MPS VII mice led to improvements in established neuropathology and restored learning and memory deficits in these animals²². In a mouse model of metachromatic leukodystrophy (MLD), Consiglio et al. demonstrated that injection of lentiviral vector carrying arylsulfatase A sequence into a single hippocampal site could establish enzyme expression capable of reducing lysosomal storage pathology and improving certain behavioral parameters²³.

Studies into treatment of the more clinically relevant MPS I have included demonstration of ex vivo retroviral gene transfer into hematopoietic cells reinfused into unirradiated adult and fetal MPS I dogs, although effectiveness of the treatment was limited by either low level gene transfer or immune response to the therapeutic gene product^24,25,26. Recently, mouse models of MPS I have also become available^27,28,29. Newborn Idua^-/- mice appear normal until about 3–5 weeks of age, when they begin to develop a characteristic flattened facial profile. By about 12–16 weeks of age, there is a broadening and thickening of the long bones as well as extensive lysosomal storage evident in Kupffer, glial, and Purkinje cells, making this animal a good model for human MPS I². Zheng et al. recently reported the use of one such MPS I mouse model to show that Idua^-/- mice receiving syngeneic bone marrow transduced with a retroviral vector carrying the human IDUA cDNA contained levels of IDUA enzyme activity that were sufficient to correct lysosomal pathology in the kidney, bladder, and fibrocartilage as well as selected regions of the brain³⁰.

We previously reported the generation of AAV vectors that transduce the human -L-iduronidase gene and correct lysosomal metabolism in fibroblasts isolated from MPS I patients³¹. Here we show that AAV-IDUA gene transfer in 1-day-old Idua^-/- mice led to high levels of plasma and tissue enzyme activities, which were sufficient to normalize urine GAG levels and reduce lysosomal storage in liver as well as cerebellar and hippocampal regions of the brain. The effect of IDUA expression on facial bone growth and CNS parameters demonstrated significant improvement of these important characteristics of MPS I. These results generally support the use of IDUA gene transfer, in particular the use of AAV-mediated gene transfer, as a therapeutic approach for MPS I.

Results

We generated the AAV vector vTRCA1 to transduce the human IDUA gene; it contains the AAV-2 inverted terminal repeats, the CMV early enhancer, the chicken -actin promoter, and the chicken -globin intron regulating expression of the human IDUA cDNA (Fig. 1A). We tested this vector first by transfection of 293 cells with pTRCA1 plasmid DNA and by transduction with packaged vTRCA1, demonstrating robust IDUA expression in both cases (data not shown).

Figure 1.

Plasma IDUA activity and urine GAG in AAV-IDUA-treated mice. (A) AAV-IDUA plasmid pTRCA1, constructed as described in Materials and Methods. TR, AAV inverted terminal repeat; CMV/BA, CMV early enhancer/chicken -actin promoter; G, chicken -globin intron; IDUA, human -L-iduronidase cDNA; pA, bovine growth hormone polyadenylation signal. (B) Plasma IDUA activity in control-treated normal and affected MPS I littermates. One-day-old mice were injected via the superficial temporal vein with 10¹⁰ particles of control AAV-gfp vector. Blood plasma was obtained as described in Materials and Methods and tested for IDUA activity at the indicated times after vector injection. () Animals 62, () 63, Idua^+/+; () 37, () 38, Idua^+/-; () 36, 39, 65, 70, Idua^-/-. (C) Plasma IDUA activity in vTRCA1-treated Idua^-/- animals. One-day-old mice were injected via the superficial temporal vein with 10¹⁰ particles of vTRCA1. Blood plasma was obtained as described in Materials and Methods and tested for IDUA activity at the indicated times after vector injection. () Animals 26, () 27, () 28, () 30, () 31, () 33, () 35, () 41, () 44, () 45. () Mean IDUA activity. (D) Urine GAG. GAG, glycosaminoglycan; cr, creatinine. Beginning 1 month after vector injection, urine was collected and GAG was quantitated as described in Materials and Methods. For Idua^-/- and Idua^+/-, graph depicts the mean (+SD) of n = 4 animals. For vTRCA1-treated Idua^-/- animals, graph depicts the mean (+SD) of n = 9 animals through month 3 and depicts the mean of n = 7 animals for months 4 and 5. ^*P < 0.05 between Idua^-/- and vTRCA1-treated Idua^-/- animals. ^**P < 0.005 between Idua^-/- and vTRCA1-treated Idua^-/- animals. Idua^+/-, untreated heterozygous controls; Idua^-/-, AAV-gfp-treated homozygous null controls.

Full figure and legend (168K)

Plasma IDUA expression, urinary GAG, and body weight measurements in vTRCA1-treated animals

To test the effects of systemic AAV-IDUA therapy on peripheral and CNS components of murine MPS I, we injected 10 1-day-old Idua^-/- mice with 1 10¹⁰ particles of vTRCA1 through the superficial temporal vein. We allowed the animals to mature for 4 weeks before undertaking any analyses of the effects of vector administration. Beginning 1 month after injection, Idua^-/- mice treated with vTRCA1 exhibited high levels of plasma enzyme activity compared to Idua^-/- animals treated with a control AAV-gfp vector (Figs. 1B and 1C). While two inadvertent animal deaths occurred during the study (animals 3728 and 3731), 6 of the 10 vTRCA1-treated Idua^-/- animals expressed levels of IDUA activity that exceeded those of control-treated Idua^+/- mice for the 5-month duration of the study. Two animals expressed levels of IDUA activity in the plasma that were at least two-fold higher than levels detected in control-treated Idua^+/+ animals throughout the study period. In addition, 2 animals continued to express heterozygote levels of enzyme activity 10 months after vector injection (mouse 30, with mean IDUA activity 8.7 3.5 nmol/mg/h, and mouse 33, with mean IDUA activity 1.8 0.4 nmol/mg/h, for months 6–10). At no time point did any control vector-treated mice exhibit detectable enzyme activity in the plasma.

Beginning at 1 month and lasting for the duration of the study, vTRCA1-treated Idua^-/- animals exhibited a significant reduction in urinary GAG compared to control-treated littermates (Fig. 1C). At each month, the mean urinary GAG level for the vTRCA1-treated group was not significantly different from levels measured in normal littermate controls. In combination with the high levels of plasma IDUA enzyme observed in vTRCA1-treated Idua^-/- animals, these results indicate a significant systemic correction of storage disease resulting from vTRCA1 administration 1 day after birth.

Weights of vTRCA1-treated and control mice determined monthly during the study assessed the effect of IDUA gene expression on growth characteristics during this period. Mean Idua^-/- control weights were higher at each time point than the mean for their control Idua^+/- littermates during the 5-month study period. The mean group weight of vTRCA1-treated Idua^-/- animals expressing detectable levels of plasma IDUA activity was normalized compared to untreated and control-treated Idua^-/- animals (Fig. 2). In the final 2 months of the study, the difference between mean vTRCA1-treated Idua^-/- weights and mean control untreated Idua^-/- animal weights was the most pronounced (P < 0.05 for months 4 and 5), further supporting the conclusion that vTRCA1 vector delivery in 1-day-old Idua^-/- mice resulted in systemic correction of the disease.

Figure 2.

Effects of AAV-IDUA treatment on Idua^-/- weights. Animals were weighed every 4 weeks beginning 1 month after vector injection. Bars depict the mean percentage difference from same sex Idua^+/- control (SEM) for n = 6 animals per group. At each time point, the heterozygous control group is defined as 100%.

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Gene transfer, IDUA expression, and lysosomal storage correction

We sacrificed five of the vTRCA1-treated Idua^-/- mice at 5 months of age for determination of IDUA gene transfer, IDUA enzyme expression, and storage materials in different organs. At this time point, four of the five vTRCA1-treated animals had plasma IDUA activity levels that ranged from approximately 3- to 10-fold higher than heterozygous control littermates (Table 1). In all five vTRCA1-treated animals studied at this time point, enzyme expression levels were highest in the heart and liver. We also detected IDUA activity in the spleen, bone marrow, and lung in the four animals demonstrating detectable plasma IDUA activity at the time of euthanization. Interestingly, of these four animals, we detected IDUA expression in the kidney in only three mice, while we detected IDUA expression in the gonad of only one animal. Three of these animals exhibited low, but detectable enzyme levels in the cerebral cortex, and two animals had low levels of gene expression in the cerebellum.

Table 1 - Tissue IDUA enzyme activity and gene copy number.

Full table

We used a real-time quantitative PCR assay developed for the human IDUA cDNA sequence³²; Pan et al., manuscript in preparation to measure the number of vTRCA1 vector copies present in different tissue samples. At 5 months, we detected gene transfer in the hearts of all five vTRCA1-treated animals ranging from approximately 0.01 to 0.94% genome equivalents (Table 1B). In three of the five animals, gene transfer in the liver ranged from 0.06 to 0.15% genome equivalents. With some exceptions, gene transfer in other tissues was undetectable (less than 0.001% genome equivalents). In all cases in which vector genomes were detected, there was also measurable IDUA enzyme (Tables 1A and 1B).

We evaluated selected tissues from the sacrificed animals for accumulation of lysosomal storage material. Gene transfer and expression in these animals were sufficient to correct lysosomal storage pathology in the liver, heart, and lung of vTRCA1-treated Idua^-/- mice (Figs. 3D, 3H, and 3L). For comparison, a liver sample from a control vector-treated Idua^-/- animal demonstrated significant lysosomal distension in resident Kupffer cells and hepatocytes (Fig. 3B). In the myocardial interstitium of the control-treated group, macrophages contained extensive pathological vacuoles (Fig. 3F). Additionally, in the lungs of this group, connective tissue between bronchioles and blood vessels contained clumps of macrophages with cytoplasm that was greatly distended by storage vacuoles (Fig. 3J). Occasional smooth muscle cells in the walls of small pulmonary arteries and bronchioles in lung sections were also noted to have vacuolar evidence of lysosomal storage material (data not shown). One vTRCA1-treated Idua^-/- animal with no IDUA activity detectable in the plasma and low levels in the heart, lung, and liver at the time of analysis (Table 2) had vacuolar pathology that was essentially indistinguishable in these regions from that of control-treated Idua^-/- animals (Figs. 3C, 3G, and 3K). These results (Table 2) demonstrate that systemic delivery of an AAV-IDUA vector can correct visceral pathology in a number of major organs of treated mice.

Figure 3.

Histopathological analysis of tissues for storage material. Group designations are as described for previous figures. One square millimeter, 1-m-thick sections were stained with toluidine blue, observed at 100 magnification, and scored for pathology as described in Materials and Methods. For liver, heart, and brain, sections from animals 35 and 27 are depicted as examples of uncorrected and corrected pathology, respectively, in the vTRCA1-treated group. In the brain, sections from animals 26 and 45 are depicted as examples of uncorrected and corrected pathology, respectively, in the vTRCA1-treated group. (A–D) Histopathology of liver. H, hepatocyte; Ku, Kupffer cell. Arrows indicate extensive cytoplasmic vacuolization of hepatocytes and Kupffer cells. (E–H) Histopathology of heart. (I–L) Histopathology of lung. (M–P) Histopathology of cerebral cortex. Arrow indicates perivascular cells distended by vacuoles. (Q–T) Histopathology of cerebellum. Arrow indicates large vacuoles in condensed Purkinje cells. (U–X) Histopathology of hippocampus. Arrow indicates perivascular cells distended by vacuoles.

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Table 2 - Summary of tissue pathology.

Full table

Vacuolization in perivascular regions of the cerebral cortex was reduced in vTRCA1-treated animals, in general, compared to control-treated mice (Fig. 3P). One AAV-IDUA-treated animal with no detectable IDUA activity in the cerebral cortex had lysosomal storage pathology that was equal in extent to that of the control-treated group (Fig. 3O). In cerebellar Purkinje cells of control-treated Idua^-/- mice, we found numerous large vacuoles that ranged from 25% of cells observed in one animal to more than 50% of cells observed (Table 2). In cerebellar Purkinje cells of vTRCA1-treated Idua^-/- animals, the presence of large vacuoles varied from nearly absent (animal 45, Table 2, Fig. 3T) to present in 10–25% of cells surveyed (animal 26, Fig. 3S). Importantly, two of the AAV-IDUA-treated animals with evidence of decreased lysosomal pathology in the cerebellum did demonstrate low but measurable IDUA enzyme activity in lysates prepared from this tissue. With the exception of one animal in this group, hippocampal perivascular vacuolization was reduced compared to control-treated Idua^-/- animals (Fig. 3X). Coupled with the results in the peripheral organs (Table 2), these assessments of pathology in the cerebral cortex, cerebellum, and hippocampus indicate that intravenous AAV-IDUA vector delivery in neonatal mice can lead to a reduced level of lysosomal storage material in the CNS as well as in peripheral tissue.

Correction of craniofacial abnormalities

One of the ultimate goals of gene therapy for treatment of MPS I and the other mucopolysaccharidoses is correction of the skeletal dysplasia associated with these diseases. To determine the effect of vTRCA1 treatment and subsequent IDUA expression on skeletal abnormalities observed in MPS I, we subjected four vTRCA1-treated Idua^-/- animals expressing the highest plasma IDUA activity and a cohort of control-treated Idua^-/- animals to computerized tomography 4 months after treatment and took precise skull width measurements. We chose three parameters of skull width in which Idua^-/- animals differed significantly (P < 0.05) from their normal littermates. In all three of these parameters, the vTRCA1-treated Idua^-/- animals as a group exhibited a trend of normalization (Fig. 4). In the width measured at the external auditory canals and the width of the zygoma at the maxilla, the observed difference was significant (P < 0.05), demonstrating that AAV transduction of the IDUA gene can improve the cranial skeletal abnormality observed in MPS I mice.

Figure 4.

Effects of AAV-IDUA treatment on craniofacial parameters. At 20 weeks after vector injection, mice were anesthetized, computed tomography was performed, and radiological distances were calculated as described in Materials and Methods. Graphs depict the means of n = 12 (SEM) for Idua^+/- and Idua^-/- control groups and n = 4 for the vTRCA1-treated Idua^-/- group. (A and C) P < 0.05 between Idua^-/- control and vTRCA1-treated Idua^-/- animals.

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Improvement of neurological function

In their severe forms, the mucopolysaccharidoses are associated with severe neurological deterioration and mental retardation. To determine the neurologic effect of AAV-mediated iduronidase gene transfer, we subjected vTRCA1- and control-treated Idua^-/- animals, as well as normal littermates, to an open field test. This test determines the effect of treatment on habituation 4 months after vector injection. Over the course of three 5-min trials for each animal, vTRCA1-treated Idua^-/- animals exhibited a trend of normalization in the percentage change in total squares covered and rearing events (Fig. 5). The vTRCA1-treated Idua^-/- group exhibited a significant reduction in the total squares covered compared to the Idua^-/- control cohort, demonstrating that AAV-IDUA vector treatment improved habituation in these mice. We conclude that AAV-mediated transduction of the IDUA gene in newborn Idua^-/- mice was sufficient to have a therapeutic impact on both systemic and neurological parameters of the disease.

Figure 5.

Open field test of habituation. At 16 weeks post-vector injection, animals were tested in three trials of 5 min per trial in the open field as described under Materials and Methods. Percentage difference describes the % change between the third and the first trial. Bars depict the means (SEM) of n = 12 for all control groups and n = 7 for the vTRCA1-treated Idua^-/- group. ^*P < 0.05 between Idua^-/- control and vTRCA1-treated Idua^-/- animals.

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Discussion

To determine the effectiveness of systemic AAV administration on a clinically relevant lysosomal storage disease, a recombinant AAV vector transducing the human -L-iduronidase gene was injected into 1-day-old MPS I mice. High levels of plasma enzyme activity were sustained for at least 5 months in 8 of 10 vTRCA1-treated Idua^-/- animals, demonstrating effective systemic transduction that was sufficient to reduce urinary GAG, clear certain tissues of lysosomal storage material, and partially correct body weight and the bony architecture of the cranium compared to control-treated Idua^-/- animals during this period. In all five vTRCA1-treated animals studied at 5 months, enzyme expression levels were highest in the heart and liver, ranging 0.5- to 17-fold above control-treated heterozygotes for heart and 0.45- to 14-fold above control-treated heterozygotes for liver. Measurable IDUA gene copy numbers that approached 1 copy per cell in the heart of one animal were achieved. However, overall levels of gene transfer measured outside the heart and liver 5 months after systemic delivery of AAV-IDUA were low, with most tissues having less than 0.001 copies per diploid genome equivalent. When subjected to an open field test, AAV-IDUA-treated animals demonstrated a statistically significant reduction in total number of squares covered and a trend of normalization in rearing events and grooming time compared to control-treated Idua^-/- mice³³.

Lysosomal storage diseases such as Gaucher disease, metachromatic leukodystrophy, and the mucopolysaccharidoses have been considered candidate disorders for treatment by gene transfer for several reasons³⁴. The concept of intercellular metabolic complementation, originally discovered by Neufeld and co-workers using cocultures of cells bearing different MPS defects⁴, has been borne out in numerous studies of cellular and genetic complementation in vitro and has been inferred from in vivo^{17,20,35,36,37,38,39} and clinical studies as well. The best clinical example of metabolic cross-correction has been the success of allogeneic bone marrow transplant in clearing peripheral storage materials in the treatment of several LSDs, ultimately mediating a significant extension of the expected life span in some individuals^9,10,11,40. Overall improvement in quality of life and life span in stably engrafted patients can be correlated directly to maintenance or improvement of cell and ultimately organ function effected by the enduring presence of active enzyme within the lysosome. Importantly, studies in Hurler children receiving bone-marrow-derived cells with normal IDUA gene sequences have demonstrated that reconstitution of systemic enzyme activity as measured in the blood leads to reduction in urinary GAG and normalization of body weight, facial features, and some skeletal manifestations of the disease^8,9,41. In addition, it has been shown in these patients that histopathological evidence of reduced lysosomal GAG storage is a reliable marker for tissue-specific recovery of active enzyme^42,43,44.

In our study, systemic enzyme activity established in MPS I mice by direct introduction of AAV-IDUA resulted in improvements in body weight and urinary GAG excretion. In addition, two of three cranial bony parameters of the disease were normalized in four of the animals expressing high levels of systemic IDUA enzyme. With the exception of one animal expressing virtually undetectable IDUA activity, vTRCA1-treated animals examined 5 months after vector injection exhibited histopathological correction of lysosomal GAG storage in the heart, lung, and liver that correlated with the presence of enzyme activity in these tissues. Delivery of normal GUSB sequences into the MPS VII mouse by bone marrow transplant or virus-mediated gene transfer have corroborated the clinical MPS I studies in establishing the link between reconstitution of systemic enzyme activity and normalization of various parameters of LSD such as GAG storage, bony changes, and body weight^{15,17,18,35,45,46}. Important studies in canine models of MPS I⁴⁷ and MPS VII^15,19 have provided additional support to the clinical studies. Recently, Zheng et al. demonstrated that following infusion of murine MPS I bone marrow cells transduced with a retrovirus carrying the human Idua cDNA, Idua^-/- recipients contained levels of active enzyme that were sufficient to reduce soluble GAG in the liver and spleen of these animals³⁰.

Stable, long-term engraftment of cells expressing normal levels of functional IDUA enzyme can lead to cognitive improvement in affected children receiving bone marrow transplant^9,48,49. In the present study, intravenous AAV-IDUA injection in neonatal MPS I mice led to an improvement in habituation behaviors compared to controls. Consistent with the overall behavioral trend in this group, some of the vTRCA1-treated Idua^-/- mice had measurable enzyme activity and most had histopathological evidence of GAG clearance in the hippocampus, a region known to be essential for habituation⁵⁰. Our data are inconclusive with respect to the source of this enzyme (i.e., whether IDUA enzyme was expressed in hippocampus or expressed elsewhere with subsequent uptake by hippocampal cells), as overall vector copy number was near background.

Direct introduction of therapeutic gene sequences into the brain of LSD mice has established a link between recovery of CNS enzyme activity, reduction of neuropathology, and normalization of cognitive deficit in these animals^21,22,23. Although there is evidence for potential cognitive benefit following intravenous delivery of normal bone-marrow-derived cells in children with Hurler syndrome, difficulties remain in finding appropriate donors, optimizing preparative regimens, and dealing with transplant procedural hindrances caused by established disease^10,11,51. Enzyme replacement in treatment of human LSD has not shown efficacy for CNS components of the disease⁵². However, in a mouse model of LSD, it has been shown that stable peripheral expression of lysosomal enzyme can impact neuropathological and neurodevelopmental hallmarks of the disease²⁰. Zheng et al. have shown that following infusion of genetically corrected murine Idua^-/- bone marrow cells, some of the syngeneic MPS I recipients contained a measurable proportion of CNS cells, including cortical neurons, choroid plexus macrophages, and cerebellar Purkinje cells, in which lysosomal storage material was reduced or in some cases eliminated³⁰. The results reported here provide evidence that perinatal AAV-IDUA delivery can lead to improvement of peripheral tissue and CNS manifestations of MPS I. Although it is possible that for larger animal models, effective clearance of lysosomal storage material in the brain may require separate, brain-directed vector targeting, introduction of corrective gene sequences in the perinatal period may represent an alternative to more invasive procedures^17,46. Indeed, it may be that compared to newborn larger animal models and newborn humans, neonatal mice are developmentally primitive and contain a relatively greater proportion of dividing stem cells, greater permeability of physiological barriers (e.g., blood–brain barrier), and a more naive immunological state⁵³. It will be important to verify the safety of AAV-mediated gene delivery in adults before initiating trials in the neonatal or in utero developmental periods in humans. Additionally, with AAV vector doses reported as high as 10¹² transducing units per kilogram in neonatal mice¹⁸, scale-up for application to large animals or humans will be technically challenging.

To date, the lack of a firm link between noninvasive, systemic delivery of a therapeutic gene and cognitive improvement in LSD prompts further evaluation. Future work will be directed toward demonstrating complete and stable cognitive improvement vis-à-vis peripheral correction of disease after systemic administration of AAV-IDUA in neonatal animals.

Materials and Methods

Construction of pTRCA1

All cloning steps were carried out according to standard methods⁴⁸. The plasmid pCAGGs and psub201 were generously provided by D. Largaespada and X. Xiao, respectively. The human IDUA cDNA and bovine growth hormone (BGH) polyadenylation signal were subcloned as an EcoRI–BglII fragment from pTRL1³¹ into pBluescript SK(-) to form plasmid subclone p1b. The CMV early enhancer/chicken -actin promoter was subcloned as an ApaI (blunt)–SspI fragment from pCAGGs into pBluescript SK(-) to form plasmid subclone pCA. The IDUA cDNA and polyadenylation sequence was excised from p1b between SacI (blunt) and EcoRI and ligated with the promoter-containing XhoI (blunt)–EcoRI fragment of pCA to yield the pCA1b subclone. An XbaI fragment containing the CMV early enhancer, -actin promoter, human IDUA cDNA, and BGH polyadenylation signal was then ligated into the XbaI site of psub201 containing the AAV terminal repeat sequences. The final plasmid, pTRCA1, was tested for IDUA gene expression in replicate cultures as described³¹.

Production and testing of rAAV-IDUA stocks

Recombinant AAV vector was packaged at the University of Minnesota according to a protocol published by Xiao et al. using the AAV-2 packaging plasmid pXX2 (generously supplied by X. Xiao)⁵⁴. Briefly, human 293 cells (American Type Culture Collection) plated at 70–80% confluence were transfected with 50 g of plasmid DNA containing equimolar ratios of pTRCA1 and pXX2. The next day, transfected cells were infected with wild-type adenovirus and approximately 72 h later, packaging cells were harvested, lysed, and AAV vector was purified using an iodixanol gradient and heparin agarose column as described⁵⁵. Vector was also produced at the University of Florida Gene Therapy Center vector facility as described⁵⁵.

All purified viral stocks were titered by dot-blot hybridization and tested for gene expression (IDUA activity) by transduction of replicate 293 cell cultures. Cytopathic effect assays in 293 cells indicated an absence of wild-type adenovirus activity in all stock AAV vector preparations. For all rAAV-IDUA stocks, the level of IDUA expressed in transduced cells was proportional to the dilution of virus used to transduce the cells. The rAAV-gfp vector vTRUF11 (generously provided by N. Muzyczka) served as control vector.

Neonatal injections

The Idua^-/- mouse colony was derived from a mating pair generously provided by E. Neufeld. Animals were maintained in a pathogen-free environment for the duration of the study. Mutant Idua^-/- animals were obtained by mating of heterozygous 4- to 8-month-old females with homozygous (-/-) 4- to 8-month-old males. Newborn pups were genotyped as described³². Twenty-four to forty-eight-hour-old pups received a 100-l injection containing 10¹⁰ particles of vTRCA1 or AAV-gfp vector vTRUF11 in lactated Ringers solution via the superficial temporal vein as described⁵⁶. Pups were left undisturbed except for weaning, cage, and feed changes for 1 month. At 5 months, animals were anesthetized with a combination of 8 mg/ml ketamine (Phoenix Pharmaceuticals, St. Louis, MO, USA), 0.1 mg/ml acepromazine maleate (Phoenix Pharmaceuticals), 0.01 mg/ml butorphanol tartrate (Fort Dodge Pharmaceuticals, Fort Dodge, IA, USA); euthanized in a CO₂ chamber; and perfused with phosphate-buffered saline, pH 7.1. Tissues were taken and divided for enzyme analyses, gene copy quantitation, and histopathology.

IDUA enzyme assays

All tissue and plasma enzyme analyses were carried out essentially as described³¹. At 1-month intervals, whole heparinized blood was collected by retroorbital eye bleeds and centrifuged to obtain plasma. Tissue samples for enzyme analyses were suspended in approximately 20 l of lysis buffer (10 mM Na₂PO₄/NaHPO₄, pH 6.0, 0.02% sodium azide, 0.1 mM dithiothreitol, 0.1% Triton X-100 per milligram of tissue)⁵⁷. Crude protein extracts were prepared from tissues either by homogenization or by three cycles of freeze–thaw using a dry ice–ethanol bath and 37°C water bath. Each tissue was processed with a separate set of surgical tools and all negative controls were processed in a separate fume hood to minimize intertissue and interanimal contamination. Lysates were cleared by centrifugation at 12,000g for 10 min at 4°C. Replicate 20-l aliquots of plasma or cleared lysate were incubated in a solution containing 2.85 mM fluorogenic substrate, 4-methylumbelliferyl--L-iduronide (Calbiochem, San Diego, CA, USA), in a 0.2 M sodium formate–0.02% Triton X buffer (pH 3.3). Sample and substrate were incubated for 6 h at 37°C before the reaction was stopped by addition of 5 ml of 0.5 M glycine buffer, pH 11. Cleaved fluorescent substrate was quantitated on a Turner fluorometer (Sunnyvale, CA, USA). Protein concentrations were determined using the Coomassie blue dye-binding assay (Bio-Rad, Hercules, CA, USA). One unit of enzyme activity is defined as the release of 1 nmol of 4-methylumbelliferyl--L-iduronide in a 1-h reaction at 37°C. Unless otherwise indicated, all results are expressed as the mean + standard deviation (SD) of replicate determinations from n = 3–8 tissue samples of animals. All statistical analyses were conducted using ANOVA, with Fisher's least significant difference algorithm as a determination of significance (P < 0.05) between groups.

GAG measurement

GAG levels were quantitated based on a method described previously⁵⁸. Urine samples were obtained one or two times per week by bladder palpation. Aliquots were diluted 1:4 and 1:10 with sodium formate buffer (pH 3.0), and replicate aliquots were mixed with 500 l 0.35 M dimethylmethylene blue dye freshly diluted in sodium formate buffer (pH 3.0). Samples were read at 535 nm and compared to a standard curve derived from heparan sulfate (Sigma, St. Louis, MO, USA) to determine the g GAG/ml. For creatinine measurement, urine samples were left undiluted or diluted 1:2 in sodium formate buffer (pH 3.0) and 5-l aliquots were mixed with 500 l freshly prepared picric acid reagent (10% saturated picric acid, 0.09 M NaOH in distilled water). Samples were measured at 535 nm on a spectrophotometer and compared to a standard curve prepared from known concentrations of creatinine (Sigma) to determine the milligrams of creatinine per milliliter. Final urine GAG values are expressed as g GAG/mg creatinine.

Real-time quantitative polymerase chain reaction (Q-PCR)

Tissue samples frozen at -20°C at the time of euthanization were carefully processed under a dissecting microscope to remove all possible connective tissue and fat. Tissues were then crudely homogenized and resuspended in lysis buffer according to the manufacturer's instructions for isolation of whole-cell DNA (Qiagen, San Diego, CA, USA). Each tissue was processed with a separate set of surgical tools and all negative controls were processed in a separate fume hood to minimize intertissue and interanimal contamination. Q-PCR for the human IDUA cDNA will be described in detail elsewhere³²; Pan et al., manuscript in preparation. Sample fluorescence signals were compared with a standard curve derived from a series of reactions using genomic DNA derived from a cell line containing a known quantity of retroviral integrant. Final values expressed as percentage genome equivalents are the means of triplicate runs.

Histopathology

Tissue samples were cut into thick sections and fixed in 2.5% glutaraldehyde in 0.1 N cacodylate buffer for at least 48 h at 4°C. Tissues were then embedded in Epon 812 resin (Electron Microscopy Sciences, Ft. Washington, PA, USA). One-square-millimeter sections, 1 m thick, were prepared and stained with toluidine blue to score for presence of pathologic storage vacuoles under blinded conditions.

Computerized tomography

Mice were anesthetized as described and computed tomography images were obtained in a Siemens Volume Zoom 4 scanner. Images were acquired in the coronal plane at 0.5-mm intervals using standard bone windowing techniques. Image analyses and measurements were done using the Image J program (http://rsb.info.nih.gov/ij/).

Open field

The open field test of habituation was performed essentially as described⁴⁹. Mice were subjected to three trials of 5 min each in a 3 3 1 ft enclosed box with a demarcated grid. Animals were divided into sex-matched control and treatment groups and scored for total squares covered (central and peripheral), number of rearing events, and grooming time per trial. Percentage differences reflect the changes from the first to the third trial in each parameter. For each mouse, trials were separated by 90 min.

References

1. Scriver, C. R. E. A. (Ed.) (2001). The Metabolic and Molecular Bases of Inherited Diseases. McGraw–Hill, New York.
2. Beighton, P. (Ed.) (1992). McKusick's Heritable Diseases of Connective Tissue, 5th ed. Mosby, St. Louis.
3. Scott, H. S., Bunge, S., Gal, A., Clarke, L. A., Morris, C. P. and Hopwood, J. J. (1995). Molecular genetics of mucopolysaccharidosis type I: diagnostic, clinical, and biological implications. Hum. Mutat. 6: 288–302. | Article | PubMed | ChemPort |
4. Fratantoni, J. C., Hall, C. W. and Neufeld, E. F. (1968). Hurler and Hunter syndromes: mutual correction of the defect in cultured fibroblasts. Science 162: 570–572. | Article | PubMed | ISI | ChemPort |
5. Kakkis, E. D., et al. (1996). Long-term and high-dose trials of enzyme replacement therapy in the canine model of mucopolysaccharidosis I. Biochem. Mol. Med. 58: 156–167. | Article | PubMed | ISI | ChemPort |
6. Kakkis, E. D., et al. (2001). Enzyme replacement therapy in feline mucopolysaccharidosis I. Mol. Genet. Metab. 72: 199–208. | Article | PubMed | ISI | ChemPort |
7. Nishioka, J., Mizushima, T. and Ono, K. (1979). Treatment of mucopolysaccharidosis: clinical and biochemical aspects of leucocyte transfusion as compared with plasma infusion in patients with Hurler's and Scheie's syndromes. Clin. Orthop. Relat. Res. 140: 194–203. | PubMed |
8. Hobbs, J. R., et al. (1981). Reversal of clinical features of Hurler's disease and biochemical improvement after treatment by bone-marrow transplantation. Lancet 2: 709–712. | Article | PubMed | ChemPort |
9. Whitley, C. B., et al. (1993). Long-term outcome of Hurler syndrome following bone marrow transplantation. Am. J. Med. Genet. 46: 209–218. | Article | PubMed | ChemPort |
10. Peters, C., et al. (1998). Hurler syndrome. II. Outcome of HLA-genotypically identical sibling and HLA-haploidentical related donor bone marrow transplantation in fifty-four children. The Storage Disease Collaborative Study Group. Blood 91: 2601–2608. | PubMed | ISI | ChemPort |
11. Peters, C., et al. (1996). Outcome of unrelated donor bone marrow transplantation in 40 children with Hurler syndrome. Blood 87: 4894–4902. | PubMed | ISI | ChemPort |
12. D'Azzo, A. (2003). Gene transfer strategies for correction of lysosomal storage disorders. Acta Haematol. 110: 71–85. | Article | PubMed | ChemPort |
13. Bosch, A. and Heard, J. M. (2003). Gene therapy for mucopolysaccharidosis. Int. Rev. Neurobiol. 55: 271–296. | PubMed | ChemPort |
14. Vogler, C., Barker, J., Sands, M. S., Levy, B., Galvin, N. and Sly, W. S. (2001). Murine mucopolysaccharidosis VII: impact of therapies on the phenotype, clinical course, and pathology in a model of a lysosomal storage disease. Pediatr. Dev. Pathol. 4: 421–433. | Article | PubMed | ChemPort |
15. Xu, L., et al. (2002). Transduction of hepatocytes after neonatal delivery of a Moloney murine leukemia virus based retroviral vector results in long-term expression of -glucuronidase in mucopolysaccharidosis VII dogs. Mol. Ther. 5: 141–153. | Article | PubMed | ISI | ChemPort |
16. Xu, L., Mango, R. L., Sands, M. S., Haskins, M. E., Ellinwood, N. M. and Ponder, K. P. (2002). Evaluation of pathological manifestations of disease in mucopolysaccharidosis VII mice after neonatal hepatic gene therapy. Mol. Ther. 6: 745–758. | Article | PubMed | ISI | ChemPort |
17. Daly, T. M., Vogler, C., Levy, B., Haskins, M. E. and Sands, M. S. (1999). Neonatal gene transfer leads to widespread correction of pathology in a murine model of lysosomal storage disease. Proc. Natl. Acad. Sci. USA 96: 2296–2300. | Article | PubMed | ChemPort |
18. Daly, T. M., Ohlemiller, K. K., Roberts, M. S., Vogler, C. A. and Sands, M. S. (2001). Prevention of systemic clinical disease in MPS VII mice following AAV-mediated neonatal gene transfer. Gene Ther. 8: 1291–1298. | Article | PubMed | ChemPort |
19. Ponder, K. P., et al. (2002). Therapeutic neonatal hepatic gene therapy in mucopolysaccharidosis VII dogs. Proc. Natl. Acad. Sci. USA 99: 13102–13107. | Article | PubMed | ChemPort |
20. O'Connor, L. H., et al. (1998). Enzyme replacement therapy for murine mucopolysaccharidosis type VII leads to improvements in behavior and auditory function. J. Clin. Invest. 101: 1394–1400. | PubMed | ChemPort |
21. Frisella, W. A., et al. (2001). Intracranial injection of recombinant adeno-associated virus improves cognitive function in a murine model of mucopolysaccharidosis type VII. Mol. Ther. 3: 351–358. | Article | PubMed | ISI | ChemPort |
22. Brooks, A. I., et al. (2002). Functional correction of established central nervous system deficits in an animal model of lysosomal storage disease with feline immunodeficiency virus-based vectors. Proc. Natl. Acad. Sci. USA 99: 6216–6221. | Article | PubMed | ChemPort |
23. Consiglio, A., et al. (2001). In vivo gene therapy of metachromatic leukodystrophy by lentiviral vectors: correction of neuropathology and protection against learning impairments in affected mice. Nat. Med. 7: 310–316. | Article | PubMed | ISI | ChemPort |
24. Lutzko, C., et al. (1999). Gene therapy for canine alpha-L-iduronidase deficiency: in utero adoptive transfer of genetically corrected hematopoietic progenitors results in engraftment but not amelioration of disease. Hum. Gene Ther. 10: 1521–1532. | Article | PubMed | ChemPort |
25. Lutzko, C., et al. (1999). Genetically corrected autologous stem cells engraft, but host immune responses limit their utility in canine alpha-L-iduronidase deficiency. Blood 93: 1895–1905. | PubMed | ISI | ChemPort |
26. Meertens, L., et al. (2002). In utero injection of alpha-L-iduronidase-carrying retrovirus in canine mucopolysaccharidosis type I: infection of multiple tissues and neonatal gene expression. Hum. Gene Ther. 13: 1809–1820. | Article | PubMed | ChemPort |
27. Clarke, L. A., et al. (1997). Murine mucopolysaccharidosis type I: targeted disruption of the murine alpha-L-iduronidase gene. Hum. Mol. Genet. 6: 503–511. | Article | PubMed | ISI | ChemPort |
28. Russell, C., et al. (1998). Murine MPS I: insights into the pathogenesis of Hurler syndrome. Clin. Genet. 53: 349–361. | PubMed | ISI | ChemPort |
29. Ohmi, K., Greenberg, D. S., Rajavel, K. S., Ryazantsev, S., Li, H. H. and Neufeld, E. F. (2003). Activated microglia in cortex of mouse models of mucopolysaccharidoses I and IIIB. Proc. Natl. Acad. Sci. USA 100: 1902–1907. | Article | PubMed | ChemPort |
30. Zheng, Y., Rozengurt, N., Ryazantsev, S., Kohn, D. B., Satake, N. and Neufeld, E. F. (2003). Treatment of the mouse model of mucopolysaccharidosis I with retrovirally transduced bone marrow. Mol. Genet. Metab. 79: 233–244. | Article | PubMed | ISI | ChemPort |
31. Hartung, S. D., Reddy, R. G., Whitley, C. B. and McIvor, R. S. (1999). Enzymatic correction and cross-correction of mucopolysaccharidosis type I fibroblasts by adeno-associated virus-mediated transduction of the alpha-L-iduronidase gene. Hum. Gene Ther. 10: 2163–2172. | Article | PubMed | ChemPort |
32. Becker, K., Pan, D. and Whitley, C. B. (1999). Real-time quantitative polymerase chain reaction to assess gene transfer. Hum. Gene Ther. 10: 2559–2566. | Article | PubMed | ChemPort |
33. Braun, S. E., Aronovich, E. L., Anderson, R. A., Crotty, P. L., McIvor, R. S. and Whitley, C. B. (1993). Metabolic correction and cross-correction of mucopolysaccharidosis type II (Hunter syndrome) by retroviral-mediated gene transfer and expression of human iduronate-2-sulfatase. Proc. Natl. Acad. Sci. USA 90: 11830–11834. | Article | PubMed | ChemPort |
34. Kohn, D. B. (1997). Gene therapy for haematopoietic and lymphoid disorders. Clin. Exp. Immunol. 107:(Suppl. 1): 54–57. | PubMed |
35. Wolfe, J. H., et al. (1992). Reversal of pathology in murine mucopolysaccharidosis type VII by somatic cell gene transfer. Nature 360: 749–753. | Article | PubMed | ISI | ChemPort |
36. Vogler, C., Sands, M. S., Levy, B., Galvin, N., Birkenmeier, E. H. and Sly, W. S. (1996). Enzyme replacement with recombinant beta-glucuronidase in murine mucopolysaccharidosis type VII: impact of therapy during the first six weeks of life on subsequent lysosomal storage, growth, and survival. Pediatr. Res. 39: 1050–1054. | PubMed | ChemPort |
37. Ohashi, T., Watabe, K., Uehara, K., Sly, W. S., Vogler, C. and Eto, Y. (1997). Adenovirus-mediated gene transfer and expression of human beta-glucuronidase gene in the liver, spleen, and central nervous system in mucopolysaccharidosis type VII mice. Proc. Natl. Acad. Sci. USA 94: 1287–1292. | Article | PubMed | ChemPort |
38. Taylor, R. M. and Wolfe, J. H. (1997). Decreased lysosomal storage in the adult MPS VII mouse brain in the vicinity of grafts of retroviral vector-corrected fibroblasts secreting high levels of beta-glucuronidase. Nat. Med. 3: 771–774. | Article | PubMed | ISI | ChemPort |
39. Kosuga, M., et al. (2000). Adenovirus-mediated gene therapy for mucopolysaccharidosis VII: involvement of cross-correction in wide-spread distribution of the gene products and long-term effects of CTLA-4 Ig coexpression. Mol. Ther. 1: 406–413. | Article | PubMed | ChemPort |
40. Krivit, W. (2002). Stem cell bone marrow transplantation in patients with metabolic storage diseases. Adv. Pediatr. 49: 359–378. | PubMed |
41. Guffon, N., Souillet, G., Maire, I., Straczek, J. and Guibaud, P. (1998). Follow-up of nine patients with Hurler syndrome after bone marrow transplantation. J. Pediatr. 133: 119–125. | PubMed | ChemPort |
42. Resnick, J. M., Whitley, C. B., Leonard, A. S., Krivit, W. and Snover, D. C. (1994). Light and electron microscopic features of the liver in mucopolysaccharidosis. Hum. Pathol. 25: 276–286. | Article | PubMed | ISI | ChemPort |
43. Resnick, J. M., et al. (1992). Pathology of the liver in mucopolysaccharidosis: light and electron microscopic assessment before and after bone marrow transplantation. Bone Marrow Transplant. 10: 273–280. | PubMed | ChemPort |
44. Braunlin, E. A., Rose, A. G., Hopwood, J. J., Candel, R. D. and Krivit, W. (2001). Coronary artery patency following long-term successful engraftment 14 years after bone marrow transplantation in the Hurler syndrome. Am. J. Cardiol. 88: 1075–1077. | Article | PubMed | ChemPort |
45. Birkenmeier, E. H., et al. (1991). Increased life span and correction of metabolic defects in murine mucopolysaccharidosis type VII after syngeneic bone marrow transplantation. Blood 78: 3081–3092. | PubMed | ISI | ChemPort |
46. Watson, G. L., et al. (1998). Treatment of lysosomal storage disease in MPS VII mice using a recombinant adeno-associated virus. Gene Ther. 5: 1642–1649. | Article | PubMed | ChemPort |
47. Shull, R. M., et al. (1987). Bone marrow transplantation in canine mucopolysaccharidosis I: effects within the central nervous system. J. Clin. Invest. 79: 435–443. | PubMed | ISI | ChemPort |
48. Peters, C., Shapiro, E. G. and Krivit, W. (1998). Hurler syndrome: past, present, and future. J. Pediatr. 133: 7–9. | PubMed | ChemPort |
49. Peters, C., Shapiro, E. G. and Krivit, W. (1998). Neuropsychological development in children with Hurler syndrome following hematopoietic stem cell transplantation. Pediatr. Transplant. 2: 250–253. | PubMed | ChemPort |
50. Cerbone, A. and Sadile, A. G. (1994). Behavioral habituation to spatial novelty: interference and noninterference studies. Neurosci. Biobehav. Rev. 18: 497–518. | Article | PubMed | ChemPort |
51. Grewal, S. S., et al. (2002). Outcome of second hematopoietic cell transplantation in Hurler syndrome. Bone Marrow Transplant. 29: 491–496. | Article | PubMed | ChemPort |
52. Desnick, R. J. and Schuchman, E. H. (2002). Enzyme replacement and enhancement therapies: lessons from lysosomal disorders. Nat. Rev. Genet. 3: 954–966. | Article | PubMed | ISI | ChemPort |
53. Stewart, P. A. and Hayakawa, E. M. (1987). Interendothelial junctional changes underlie the developmental 'tightening' of the blood–brain barrier. Brain Res. 429: 271–281. | PubMed | ChemPort |
54. Xiao, X., Snyder, R. O. and Samulski, R. J. (1996). Production of recombinant adeno-associated viral vectors. Current Protocols in Human Genetics, (pp. 12.11–12.23). Wiley: New York.
55. Zolotukhin, S., et al. (1999). Recombinant adeno-associated virus purification using novel methods improves infectious titer and yield. Gene Ther. 6: 973–985. | Article | PubMed | ISI | ChemPort |
56. Sands, M. S. and Barker, J. E. (1999). Percutaneous intravenous injection in neonatal mice. Lab. Anim. Sci. 49: 328–330. | PubMed | ChemPort |
57. Huang, M. M., Wong, A., Yu, X., Kakkis, E. and Kohn, D. B. (1997). Retrovirus-mediated transfer of the human alpha-L-iduronidase cDNA into human hematopoietic progenitor cells leads to correction in trans of Hurler fibroblasts. Gene Ther. 4: 1150–1159. | Article | PubMed | ChemPort |
58. Whitley, C. B., Draper, K. A., Dutton, C. M., Brown, P. A., Severson, S. L. and France, L. A. (1989). Diagnostic test for mucopolysaccharidosis. II. Rapid quantification of glycosaminoglycan in urine samples collected on a paper matrix. Clin. Chem. 35: 2074–2081. | PubMed | ChemPort |

Acknowledgements

We thank D. Largaespada for the plasmid pCAGGS and the participants in the Gene Therapy for Metabolic Disorders Program Project Grant for critical input and technical advice. This work was supported by Grant HD32652 from the National Institutes of Health.