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J. Biol. Chem., Vol. 278, Issue 35, 33501-33518, August 29, 2003
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Mice Lacking Phosphatidylinositol Transfer Protein-
Exhibit Spinocerebellar Degeneration, Intestinal and Hepatic Steatosis, and Hypoglycemia*
¶ 
From the
Department of Cell and Developmental
Biology, Lineberger Comprehensive Cancer Center, University of North Carolina,
Chapel Hill, North Carolina 27599-7090, the
Department of Neurology, University of Michigan
School of Medicine, Ann Arbor Veterans Affairs Medical Center GRECC, Ann
Arbor, Michigan 48104-0520, the ||Department of
Nutrition, University of Alabama at Birmingham, Birmingham, Alabama 35294, and
**University of California School of Medicine, San
Diego, La Jolla, California 92093-0644
Received for publication, April 7, 2003 , and in revised form, May 30, 2003.
 |
ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Phosphatidylinositol transfer proteins (PITPs) regulate the interface between lipid metabolism and cellular functions. We now report that ablation of PITP
function leads to aponecrotic spinocerebellar disease,
hypoglycemia, and intestinal and hepatic steatosis in mice. The data indicate
that hypoglycemia is in part associated with reduced proglucagon gene
expression and glycogenolysis that result from pancreatic islet cell defects.
The intestinal and hepatic steatosis results from the intracellular
accumulation of neutral lipid and free fatty acid mass in these organs and
suggests defective trafficking of triglycerides and diacylglycerols from the
endoplasmic reticulum. We propose that deranged intestinal and hepatic lipid
metabolism and defective proglucagon gene expression contribute to
hypoglycemia in
PITP–/–
mice, and that hypoglycemia is a significant contributing factor in the onset
of spinocerebellar disease. Taken together, the data suggest an unanticipated
role for PITP in with glucose homeostasis and in mammalian endoplasmic
reticulum functions that interface with transport of specific luminal lipid
cargoes. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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PITPs1 mobilize PtdIns or PtdCho between membrane bilayers in vitro (1, 2). In vivo studies demonstrate that PITPs control the interface between membrane trafficking and lipid metabolic pathways in yeast (3–6). By contrast, the physiological functions for mammalian PITPs, which are structurally unrelated to yeast PITPs (7, 8), are not understood at either the cellular or organismal levels.
Mammals express at least three soluble PITPs: PITP, PITP
, and
rdgB
(9–11).
PITP and PITP share 77% primary sequence identity, are encoded by
distinct genes, and exhibit biochemical differences. Yet both PITP and
PITP (and even yeast PITPs) function as soluble factors that stimulate
various reconstitutions of PIP-dependent functions in permeabilized mammalian
cells. These functions include regulated and constitutive membrane trafficking
and phospholipase C-dependent signaling through G-protein-coupled receptors
(12–14).
Given the lack of PITP specificity in these assays, it remains unclear how
faithful such reconstitutions are in reporting physiological functions for
mammalian PITPs.
Genetic studies are providing initial clues regarding PITP function in
metazoans. An inherited form of light-enhanced retinal degeneration in
Drosophila results from inactivation of a membrane-bound PITP
(15). In mice, reduction of
PITP to 18% of wild-type levels is the basis for the vibrator
neurodegenerative disorder
(16,
26). Gene ablation approaches
suggest PITP plays an essential housekeeping function, whereas
PITP is nonessential for ES cell viability and is not a quantitatively
significant factor in membrane trafficking, PIP metabolism, or growth factor
signaling in ES cells
(17).
In this report, we describe the consequences associated with ablation of
PITP function in the mouse. We find that PITP, although
dispensable for prenatal development, is required for neonatal survival.
PITP–/–
neonates suffer from a severe spinocerebellar neurodegenerative disease and
exhibit defects in dietary fat and -tocopherol transport across the
small intestine. This intestinal steatosis in some respects resembles CRD, a
human disorder of unknown molecular etiology
(18,
19). Liver steatosis is also
prominent in the mutant mice, suggesting the possibility that PITP
nullizygosity also compromises lipoprotein assembly and/or neutral lipid
secretion in hepatocytes. Finally,
PITP–/–
mice are severely hypoglycemic. Our results suggest a novel and unanticipated
role for PITP in regulating cargo-specific lipid transport from the
enterocyte and hepatocyte ER, endocrine pancreas function, and glycogen
metabolism.
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Generation and Genotyping of PITP
–/–
Mice—AB1-derived
+/PITP
1::neo* ES cells have been
described previously (17). The
+/PITP::neo/puro ES cells corresponded to
the OST 1152 line (Lexicon Genetics OmniBankTM library
(20)). Mice were generated by
injection of ES cells into C57BL/6 blastocysts, implantation of blastocysts
into pseudopregnant foster mothers, and identification of male chimeric mice
competent for germ line transmission of each allele. Genotypes for
PITP1::neo* mice (JG line) were
determined by using a three-primer PCR assay. We employed a primer specific
for the homozygous
PITP+/+ genotype
(AB-2; 5'-GCGAGGCATCACTCTTCCCCTC-3'), the heterozygous
PITP–/+
genotype (AB-1B; 5'-CACCATCCCCCACGGTGACTG-3'), and the
PITP–/–
genotype (PG-1; 5'-GAATGTGTGCGAGGCCAGAGG-3') in a 33-cycle
reaction (53 °C annealing temperature). Genotypes for
PITP::neo/puro mice (L1 line) were determined in two
steps. First, a two primer assay that monitored Neo distinguishes
PITP+/+ from
PITP–/+ and
PITP–/–
mice. Primers GE-UP (5'-GGGCGCCCGGTTCTTT TTGTGA-3') and GE-DO
(5'-TTGGTGGTCGAATGGGCAGGTAGC-3') were used in a 28-cycle reaction
(60 °C annealing temperature). To distinguish
PITP–/+ from
PITP–/–
L1 mice, we resorted to immunoblot analyses of mouse brain using
PITP-specific serum
(17).
Serum Analyses—Blood was collected from mice immediately
after heart puncture and clotted, and serum was clarified by centrifugation.
Serum glucose was determined by using either the Trinder assay (Sigma) or was
measured by Antech Diagnostics (Farmingdale, NY). Insulin and
-hydroxybutyrate were determined using the Immunoassay System (Crystal
Chem Inc.) and the -hydroxybutyrate assay kit (Sigma), respectively. All
other serum analyses were performed by Antech Diagnostics (Farmingdale,
NY).
Carcass Analyses—Carcass analyses were as described previously (21). Gastrointestinal tracts were removed (stomach, small and large intestine, and cecum) and carcasses weighed. Body water content of eviscerated carcasses was determined by drying to constant weight in a 60 °C oven and measuring differences between the pre- and post-drying carcass mass. Dried carcasses were minced, ground to a homogeneous mixture, and extracted with petroleum ether in a Soxhlet apparatus to determine fat mass and fat-free dry mass. Fat-free dry mass was burned overnight at 600 °C(>8 h) to determine eviscerated carcass ash.
Histological Analysis—Mice were anesthetized with 1.25% Avertin and perfused with phosphate-buffered saline, 4% paraformaldehyde. Duodenum, ileum, cerebellum, pancreas, and spinal cord were harvested, flushed with fixative (duodenum and ileum only), dissected, and infused with fixative for 24 h. Samples were mounted in paraffin, and 5-µm-thick sections were stained and mounted. These sections were rehydrated by serial transfer from xylene to 50% EtOH, stained with hematoxylin/eosin, and mounted in Permount (Fisher). Where osmium staining was employed, duodenal and liver sections were stained in 5% potassium dichromate, 2% osmium tetroxide for 8 h prior to paraffin embedding (22) and counterstained with toluidine blue O.
Whole brains were extracted from mice perfused with 4% paraformaldehyde and 2.5% glutaraldehyde, washed for several days in phosphate-buffered saline, and mounted in paraffin. Brains were sliced in half along the sagittal plane; each half was embedded in paraffin; and 5- or 8-µm-thick slices were mounted onto treated slides. Sections were rehydrated to 50% EtOH. For visualization of Purkinje cells, slices were incubated with goat anti-calbindin antibodies (Santa Cruz Biotechnology; Santa Cruz, CA) and developed with the Vectastain ABC kit (Vector Laboratories, Burlingame, CA). After incubation in 2% osmium fumes for 10 min, slices were counterstained with toluidine blue O.
For Oil Red O staining, livers were extracted from mice perfused with 4% paraformaldehyde, washed with phosphate-buffered saline, and frozen at –20 °C. Frozen livers were mounted, sectioned (8 µM), fixed to a histological slide, and placed in absolute propylene glycol (2 min). Slides were moved into Oil Red O solution (Newcomer Supply, Middleton, WI, catalog number 12722) for 1 h, differentiated in 85% propylene glycol (1 min), rinsed 2x in distilled water, counterstained with hematoxylin (10 s), and mounted in glycerin.
Electron Microscopy—Mice were perfused with 4% formaldehyde, 2.5% glutaraldehyde. Biopsies from intestine, liver, and spinal cord were post-fixed with 1% OsO4, dehydrated with acetone, embedded in epoxy (23), sectioned (65 nm-thick), and stained with 4% uranyl acetate and Sato's lead mixture (24). Samples were viewed at 80 kV in a Phillips Tecnai 12 microscope (FEI Co., Eindhoven, The Netherlands) and imaged with a Gatan MultiScan model 794 digital camera (Gatan, Pleasanton, CA).
Epoxy-embedded samples were sectioned for histological analysis (2-µm thickness) and stained with 1% toluidine blue O in 1% sodium borate. Digital images were collected with a SPOT RT digital camera (Diagnostic Instruments, Inc., Sterling Heights, MI), using Plan Fluor Nikon objectives mounted in a Nikon Eclipse E400 microscope (Nikon Inc., Melville, NY). Morphometric analysis of spinal cord sections was carried out with Scion Image software (Scion Corp., Frederick, MD).
-Tocopherol Analyses—Brain -tocopherol was
extracted as described (25),
injected onto a 250 x 4.6 mm Phenomenex ODS 20 reversed phase C18 column
(5-µm particle diameter), and eluted with methanolic 0.05% ammonium acetate
using a flow rate of 1.5 ml/min. The high pressure liquid chromatography
system consisted of a PerkinElmer Life Sciences model LC200 gradient pump, an
AS 200 Autosampler, and an LC 295 programmable UV-visible light detector set
at 292 nm.
Liver Glycogen Analysis—Quantification of liver glycogen was by the method of Passonneau and Lauderdale (26). Glycogen was extracted from acidified liver homogenates and hydrolyzed to glucose with amyloglucosidase (Sigma). Glucose was determined by a glucose oxidase-coupled Trinder assay (Sigma).
Adenylate Nucleotide Analysis—ATP and ATP/ADP ratios were measured using the ApoGlowTM kit (BioWhittaker Molecular Applications; Rockville, MD). Samples (0.5 mg tissue) were collected and rapidly frozen. Tissues were thawed and homogenized in the presence of a nucleotide-releasing mixture supplied by the manufacturer. Acid-extracted ATP was assayed with a luciferase-based system with picomolar sensitivity. ADP was converted to ATP and measured after a 5-min incubation at 22 °C.
Pancreatic Histology—Whole pancreas from mice perfused with 4% paraformaldehyde was embedded in paraffin and serially sectioned (thickness = 5 µm). Islet numbers were assessed by sequential observation of hematoxylin/eosin-stained sections. Care was exercised to separate islets from patches of connecting ducts and intervening connective tissue, and not to re-score the same islet in successive sections. Islet-like, encapsulated structures larger than 100 µm that were detected in at least three consecutive sections were scored as islets. The total pancreatic area studied was similar in all sections.
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RESULTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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PITP
–/–
Mice Develop to Term—Ablation of PITP function in the
mouse was achieved using two independent null alleles. First, a homologous
recombination vector was constructed where exons 8–10 of the PITP
gene are replaced with a neo cassette
(Fig. 1A). This
mutation (PITP::neo*) deletes PITP
residues 162–257, a region critical for PITP function
(17). Second, survey of the
Lexicon Genetics OmniBankTM gene trap library (see "Experimental
Procedures") identified an insertion mutation in the PITP
structural gene (PITP::neo/puro). This
allele is genetically similar (although not identical) to
PITP::neo* as it also truncates PITP
after residue 162 (Fig.
1B). Mice were derived from each targeted ES cell line.
PITP–/+
offspring are phenotypically normal and fertile, and mice homozygous for
either of these two mutations exhibit indistinguishable phenotypes. The
phenotypic data presented below were obtained from both
PITP1::neo* and
PITP::neo/puro homozygous animals.
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Intercrosses with heterozygous mice carrying either
PITP1::neo* or the
PITP::neo/puro allele yielded genotypes
consistent with a fully penetrant autosomal recessive mutation. From a
dedicated pool of 408 live births, 89
PITP–/–
progeny were recovered (Fig.
1C). The genotypic distribution of 106
PITP+/+, 213
PITP–/+, 89
PITP–/–
corresponds to a 1.000:2.009:0.840 ratio that approximates closely the 1:2:1
ratio predicted by Mendel's rules. Correct gene targeting in the progeny was
verified by PCR genotyping and immunoblotting of brain extracts with a
specific PITP antiserum (Fig.
1D). Antibodies directed against the PITP N
terminus failed to detect a truncation product in mice homozygous for
PITP1::neo* or
PITP::neo/puro, suggesting that both alleles
represent null mutations. Finally, we find that relative PITP levels are
unchanged in PITP+/+
and
PITP–/–
brain (Fig. 1D),
indicating that
PITP–/–
mice do not activate compensatory processes that increase PITP
expression.
Neonatal Mortality of
PITP–/–
Mice—PITP–/–
mice failed to thrive and died at a very young age. In a sample pool of 57
PITP–/–
mice, 40% died within 48 h after birth
(Fig. 2A). These early
P0 and P1 deaths were not characterized by obvious external abnormalities.
Moreover, postmortem analyses revealed that the stomachs of the expired
PITP–/–
mice contained copious quantities of milk, indicating that mutant animals had
nursed. Of the
PITP–/–
progeny that survived past P1, a steady incidence of mortality was observed
between P2 and P11. Almost all mice expired by P11, and only one
PITP–/–
mouse lived to P14 (Fig.
2A).
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PITP–/–
progeny that survived past P1 were initially indistinguishable from
PITP+/+ and
PITP–/+
littermates in size, external morphology, and behavior. By P4, however, two
phenotypes rapidly asserted themselves. First, most
PITP–/–
mice were moribund and exhibited little spontaneous movement. These mice did
respond to touch, however. By contrast, 
10% of the
PITP–/–
mice experienced spontaneous seizures. All
PITP–/–
mice were severely ataxic and were generally incapable of maintaining
themselves upright. We also observed coarse action tremors upon limb extension
in
PITP–/–
mice. Second, these mutant mice failed to thrive. Although the
PITP–/–
mice gained body mass, they did so slowly
(Fig. 2B). By P10,
surviving
PITP–/–
mice were 2.5-fold less massive than their
PITP+/+ and
PITP–/+
littermates (Fig.
2C).
PITP–/–
Mice Suckle Effectively—Several lines of evidence indicate that
the failure of
PITP–/–
mice to thrive is not the simple consequence of neurological defects. We
observed
PITP–/–
mice in the act of suckling, and postmortem analyses indicated both copious
quantities of milk in stomachs of these mice
(Fig. 2D) and
substantial amounts of digested matter throughout the
PITP–/–
intestinal tract (not shown). The suckling competence of
PITP–/–
mice notwithstanding, the mutant animals exhibited pathologically low body fat
levels. This reduction is obvious when the subcutaneous fat pads of
PITP+/+ and
PITP–/–
mice are compared. Whereas
PITP+/+ controls
exhibit large axillary and inguinal fat pads, these structures are absent from
PITP–/–
animals (Fig. 2D).
Chemical analyses of eviscerated carcasses quantified these differences;
PITP+/+ and
PITP–/–
mice exhibited total body fat contents of 15.0 ± 1.1 and 4.0 ±
0.5% of total body mass, respectively (Fig.
2E). Significantly, the relative water contents of
PITP+/+ and
PITP–/–
carcasses are similar (Fig.
2E), indicating that
PITP–/–
mice are not dehydrated, as would be expected if there were substantial
suckling defects. The relative fat-free dry mass contents and carcass bone ash
contents are also comparable (Fig.
2E). Consistent with those measurements, we find that
organ/total body mass ratios in
PITP–/–
mice for brain, liver, and other major organs are also proportional to total
body mass (not shown).
Elevated Apoptosis and Purkinje Cell Defects in
PITP–/–
Cerebellum—Nissl staining does not reveal obvious defects in
development or morphology of
PITP–/–
cerebrum, thalamus, hippocampus, or cerebellum. Normal cellularity was
observed in all regions examined (data not shown). However, PITP
insufficiencies evoked functional defects in the cerebellum. These defects
were apparent at several levels. Whereas TUNEL staining showed a sparse and
random distribution of apoptotic foci in wild-type brain, apoptosis was more
prevalent throughout
PITP–/–
cerebellum. This is particularly evident in the external granule layer of
mutant cerebellum (Fig.
3A).
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Second, we find Purkinje cell defects in
PITP–/–
animals. Purkinje cells normally align themselves into a sharply defined
monolayer that lies between the molecular and external granular layers of the
cerebellum (Fig. 3B).
These cells elaborate well developed apical dendritic stalks which arborize
into luxurious branches that penetrate into the molecular layer of the
cerebellum.
PITP–/–
Purkinje cells, while retaining normal flask-shaped cell body morphologies,
exhibit either abbreviated apical dendritic stalks or no obvious stalks at all
(Fig. 3B). Moreover,
the dendritic branches emanating from the abbreviated apical stalks are less
arborized than those of
PITP+/+ Purkinje
cells. We also observed cases where these cells exhibit defects in spatial
alignment, reside off of the defined layer, or are otherwise misoriented.
Degenerative Disease in
PITP–/–
Cerebellum—PITP–/–
cerebellum suffered major degenerative insult as judged by the extent of
reactive gliosis. Wild-type cerebellum shows low levels of GFAP, a specific
marker for activated astrocytes (Fig.
3C). By contrast,
PITP–/–
cerebellum exhibits a dramatic staining for GFAP, and reactive gliosis is
especially prominent in the white matter trunk of this organ
(Fig. 3C). Even in
these relatively less affected areas, GFAP immunohistochemistry suggests some
10% of the cells represent activated astrocytes that are frequently seen to be
enveloping neuron cell bodies (Fig.
3D). Reactive gliosis is not observed throughout the
mutant brain. Whereas
PITP–/–
cerebellum and brain stem are heavily infiltrated with activated glial cells,
and the subthalamic region is also involved, significant gliosis is not
observed in the neocortex, striatum, hippocampal formation, and other
forebrain regions (not shown).
Finally, electron microscopy reveals ER defects in cerebellar neurons of
PITP–/–
mice. These defects manifest themselves as significant vacuolations of the
smooth ER region, even though adjacent regions of rough ER retain normal
morphology (not shown). Such vacuolations are not observed in smooth ER of
PITP+/+ cerebellar
neurons.
Neurodegeneration and Inflammation in
PITP–/–
Spinal Cord—We consistently observed WM deficits in
PITP–/–
cervical, thoracic, and lumbar spinal cord. Whereas ventral WM is well
developed in PITP+/+
cervical spinal cord, the corresponding WM regions are thin and sparse in
PITP–/–
animals (Fig. 4A). WM
to GM area ratios in cervical spinal cord are 0.45 ± 0.1 for
PITP+/+ and 0.32
± 0.02 for
PITP–/–
mice, respectively (p < 0.01; n = 6). In addition to the
WM deficits, evidence of inflammation pervades all regions of the
PITPa–/– spinal
cord. Toluidine blue O-staining reveals areas of abnormally heavy staining in
the ventral horn where motor neuron cell bodies reside, suggesting cell damage
in these areas (Fig.
4A). Moreover, whereas the WM/GM interface of
PITP+/+ cervical
spinal cord presents normal vascular structures, neurons, and accessory cells,
corresponding regions of the
PITP–/–
spinal cord exhibit densely stained cells and damaged neurons
(Fig. 4B). These
pathologies encompass a spectrum of cell death events. These range from
occasional apoptotic neurons (condensed nuclei, fragmented cytoplasm; not
shown) to large numbers of neurons undergoing aponecrotic processes
(Fig. 4B). Aponecrosis
is a form of cell death associated with reductions in cellular ATP and
increases in ADP levels (27,
28). Analyses from six
PITP–/–
mice indicate that 18 ± 1% of the total motor neuron cell bodies in the
ventral horn exhibit properties of aponecrosis. These include pericytoplasmic
vacuolation, reduced cytoplasmic contents, and cytoplasmic proliferation of
irregular electron-translucent vesicles
(Fig. 4B).
|
We did not observe aponecrotic motor neurons in the ventral horn of
PITP+/+ spinal cord.
Extensive vacuolation and membrane blebbing is prevalent in cells that line
the vasculature of
PITP–/–
spinal cord (not shown). Axons surrounding capillaries and small vessels in
the GM are enlarged and often damaged. This is unlikely to represent a
perfusion artifact, as this phenotype was not recorded in any of the spinal
cord sections analyzed from 14
PITP+/+ mice.
Toluidine blue O-staining reveals the presence of cells containing an
abundance of purple intracellular granules in the perivascular matrix
(Fig. 4C, left
panel). This obvious purple metachromasia is diagnostic of inflammatory
mast cells (29,
30). Accordingly, we find
extratissular macrophages in perivascular tissue or even in the vessels
themselves (Fig. 4C,
right panel), suggesting the blood/brain barrier of
PITP–/–
mice is itself compromised. Finally, spinocerebellar injury in
PITP–/–
mice also includes processes that resemble those of other myelin-related
central nervous system inflammatory disorders
(31). First, macroscopic
swelling in the dorsal spinal columns is apparent. The non-neuronal area
(neuropil) comprises 54 ± 3% in
PITP+/+ spinal cord
(n = 6; p = 0.01) and 70 ± 6% of total area in
PITP–/–
spinal cord, respectively. Degenerative processes in neuropil, typified by
swollen axons lacking organelles and cytoskeletal filaments, are also obvious
(not shown). Second, demyelination is scored in both WM and GM areas of
PITP–/–
mice and especially in the dorsal spinal columns. Supporting cells that may be
remodeling myelin are also observed (Fig.
4D).
The fragility of the mutant central nervous system notwithstanding, a
variety of
PITP–/–
neurons are amenable to primary culture. Cultured cortical neurons, cerebellar
granule cells, and spinal cord dorsal root ganglion neurons from
PITP–/–
mice are not more fragile than their
PITP+/+ counterparts
in any obvious way. Indeed, titration of nerve growth factor concentrations
from 50 to 5 ng/ml revealed no differences in the thresholds of trophic factor
required to sustain viability of
PITP–/–
versus PITP+/+
dorsal root ganglion neurons in culture (not shown).
PITP–/–
cerebellar granule cells are similarly robust. These data suggest a
significant cause of neuronal injury in
PITP–/–
mice is a hostile physiological environment, rather than some overriding cell
autonomous defect.
Lipid Dysregulation in
PITP–/–
Enterocytes—Failure of
PITP–/–
mice to thrive suggests a malabsorption disorder. Histological analyses of
duodenum indicate the architecture of this organ and its muscle layers are
similar in PITP+/+
and
PITP–/–
mice. Moreover, intestinal motility also appears normal (not shown). There are
obvious anomalies in
PITP–/–
intestinal epithelium, however. Most strikingly,
PITP–/–
enterocytes exhibit an obviously heavy impregnation with the lipophilic agent
osmium relative to
PITP+/+ enterocytes
(Fig. 5A). These
histological properties are consistent with an abnormal accumulation of
intracellular lipid in mutant enterocytes. If the accumulated lipid is derived
from dietary fat, rather than manifestation of some other cellular pathology,
active feeding is expected to be required for the loading of mutant
enterocytes with lipid. Indeed, enterocytes harvested from
PITP–/–
mice subjected to a 10-h fast are clear of accumulated lipid (not shown).
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Closer inspection of
PITP+/+ duodenal
enterocytes by electron microscopy demonstrates that these cells contain a
heterogeneous population of lipid bodies
(Fig. 5B). Even at
this high resolution, these structures stain with osmium in a manner
consistent with a lipid interior, and these vesicles exhibit an average
diameter of 300–700 nm. Inspection of the borders of these lipid bodies
at very high magnifications reveals a classical trilaminar membrane
surrounding these structures (not shown).
PITP–/–
enterocytes, on the other hand, are packed with vesicles that are also
membrane-enclosed but are much larger than those observed in
PITP+/+ enterocytes.
Moreover, the structures that accumulate in
PITP–/–
enterocytes often appear multilobed (Fig.
5C). Representative profiles of such structures from
PITP+/+ and
PITP–/–
enterocytes are shown at higher magnification in
Fig. 5D. The size
differences of the structures in wild-type and nullizygous enterocytes
notwithstanding, these share many structural features. It is therefore likely
that the biogenesis of the lipid bodies initially follows a common course in
PITP+/+ and
PITP–/–
enterocytes.
Morphometric measurements indicate a rather unimodal frequency distribution
of vesicle perimeters from
PITP+/+ mice
(average is 2 µm; Fig.
5E). By contrast, the accumulated structures in
PITP–/–
enterocytes are much larger and have multimodal size distributions. Perimeters
of 20 µm are often observed, and perimeters in excess of 50 µm are also
encountered (Fig. 5E).
Finally, our results indicate that the lipid structures in
PITP–/–
enterocytes reside predominantly in dilated regions of smooth ER
(Fig. 5F).
-Tocopherol and Plasma TG Deficiency in
PITP–/–
Mice— The failure of
PITP–/–
mice to thrive, when coupled with their retention of lipid in the enterocyte
smooth ER, predicts a malabsorption-mediated deficit in fat-soluble vitamins,
particularly -tocopherol (vitamin E), and reduced levels of
post-prandial plasma TGs. We find
PITP–/–
mice are deficient in -tocopherol. Whereas
PITP+/+ brain
exhibits 19.48 ± 3.41 µg of -tocopherol/gm tissue, values for
PITP –/–
brain are 10-fold reduced (2.11 ± 0.04 µg of
-tocopherol/g of tissue; Fig.
5G). Post-prandial plasma TG levels in
PITP–/–
mice are also 3-fold reduced relative to
PITP+/+ mice (55
± 13 versus 183 ± 15 mg/dl).
By contrast,
PITP+/+ and
PITP–/–
plasma cholesterol levels are similar. Indeed, comprehensive blood chemistry
analyses demonstrate many (although not all) parameters are largely normal for
PITP–/–
mice (Table I). Immunoblot
experiments indicate that apolipoprotein B48 and microsomal triglyceride
transfer protein, two components required for chylomicron assembly, are
expressed in
PITP–/–
mice and that serum apoB levels are similar to those recorded for
PITP+/+ mice (not
shown).
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PITP–/–
Mice Exhibit Microvesicular Steatosis in Liver— The pathways
for lipid transport from the ER lumen to the cell surface are similar in
enterocytes and hepatocytes. In that regard,
PITP–/–
liver also exhibits obvious microvesicular steatosis, as evidenced by its
dramatically enhanced staining with osmium
(Fig. 6A) and with the
lipophilic dye Oil Red O (Fig.
6B). Because Oil Red O exhibits specific affinity for
neutral lipids (e.g. TG, DAG, and CE), we conclude that the
accumulated lipid is largely of the neutral class. These histochemical data
are extended by electron microscopy. Relative to wildtype hepatocytes,
PITP–/–
cells exhibit an abundance of large intracellular lipid bodies. We often
detect such structures in the nuclear matrix of mutant hepatocytes, a
circumstance we have yet to encounter in wild-type cells
(Fig. 6C).
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The lipid bodies are either membrane-bound or represent large cytoplasmic
lipid droplets (Fig.
6D). Unlike in mutant enterocytes, where accumulated
lipid is predominantly found in membrane-enclosed compartments, 60% of
the lipid bodies in
PITP–/–
hepatocytes represent what we interpret to be cytosolic lipid droplets due to
lack of identifiable boundary membrane
(Fig. 6D). This raises
the possibility that the mechanism of hepatic steatosis differs from that of
the
PITP–/–
enterocytes. Although we cannot yet exclude the possibility that allocation of
fatty acids and lipids between storage pools and lipoprotein pools is deranged
in
PITP–/–
liver, at least some key fatty acid and lipid biosynthetic enzymes (acetyl-CoA
carboxylase, stearyl-CoA reductase, and fatty-acid synthase) are not
transcriptionally up-regulated in this mutant tissue (not shown). Moreover,
the hepatic microvesicular steatosis, although qualitatively resistant to a
short 6-h fast, is reversed by fasting mutant mice for 17 h (not shown).
Quantitative Analyses of Lipids and Fatty Acids in
PITP–/–
Liver and Brain—The lipid derangement in
PITP–/– duodenum and liver suggests increased
neutral lipid mass. To test this interpretation directly, the mass of a
variety of lipid species in
PITP+/+ and
PITP–/–
liver and brain was quantified. The intestine provides technical challenges in
this assay as the mice analyzed are neither fasted nor germ-free. We assume
analysis of liver lipid mass is generally representative of the situation in
intestine.
As shown in Fig.
7A, although the relative TG, DAG, and CE contents are
very similar in matched wild-type and PITP nullizygous brain tissue,
bulk neutral lipid mass is dramatically elevated in PITP nullizygous
liver. Bulk TG and FFA mass is 2.0- and 2.6-fold increased relative to the
respective values recorded for wild-type liver. Bulk CE and DAG mass is also
increased in
PITP–/–
liver. Unexpectedly,
PITP–/–
brain exhibits a 3.3-fold increase in FFA mass relative to
PITP+/+ brain
(Fig. 7A).
|
The derangements in neutral lipid/FFA content in
PITP–/–
tissues do not reflect a universal derangement of lipid homeostasis. PtdCho,
phosphatidylethanolamine, and phosphatidylserine mass values are similar in
matched wild-type and PITP nullizygous brain and liver tissue
(Fig. 7B). We do note,
however, that mass of specific polar lipids such as sphingomyelin and
lyso-PtdCho are elevated in PITP nullizygous brain, and CL levels are
altered in both mutant brain and liver relative to matched wild-type tissues
(Fig. 7C).
Fatty acid compositions of the CE, DAG, TG, FFA, sphingomyelin, and CL
pools of PITP+/+ and
PITP–/–
brain (Table II) and liver
(Table III) were also compared.
The most striking qualitative and quantitative difference is the reduced
content of 18:2n6 (9,12-octadecadienoic acid) in mutant brain TG,
DAG, and CL pools (Table II),
and an increase in 18:1n9 (9-octadenoic acid) in CE, DAG, and FFA
pools in nullizygous liver (Table
III). Fatty acid profiles of brain and liver lyso-PtdCho were
determined for
PITP+/+ and
PITP–/–
mice, and no differences were scored (not shown).
|
|
Reduced ATP Content in
PITP–/–
Cerebellum and Liver— Electron microscopic analyses reveal not
only lipid accumulation but also indications of aponecrosis in mutant
hepatocytes. These indications resemble those observed in the
PITP–/–
spinocerebellar system (see above) and include the significantly increased
proliferation of electron-dense autophagic bodies throughout the cytoplasm
(cytolysosomes) and regions of dilated smooth ER (not shown). Aponecrosis in
the spinocerebellar system and in liver suggests a fundamental defect in
energy metabolism in PITP nullizygous mice. Indeed, ATP levels are
diminished in
PITP–/–
relative to PITP+/+
liver (40.4 ± 8.6 versus 98.1 ± 9.3 pmol of ATP/mg of
tissue; Fig. 8A).
ATP/ADP ratios further emphasize this reduction (2.8 ± 0.5 for
PITP–/–
and 9.1 ± 3.1 for
PITP+/+ liver;
Fig. 8B). Similarly,
PITP+/+ and
PITP–/–
animals exhibit 83.3 ± 5.3 and 44.9 ± 5.9 pmol of ATP/mg of
cerebellum, respectively (Fig.
8A). Reduced cerebellar ATP/ADP ratios are measured as
well (7.7 ± 1.0 for wild-type versus 4.2 ± 0.5 for
mutant; Fig. 8B).
Interestingly,
PITP+/+ and
PITP–/–
brain ATP levels (40 ± 59 and 343 ± 65 pmol of ATP/mg of brain,
respectively) and ATP/ADP ratios (13.6 ± 2.3 versus 9.1
± 0.7, respectively) are comparable, thereby regionally correlating
neuronal and hepatic injury with reduced cellular ATP
(Fig. 8, A and
B). The significant decrease in mass of the mitochondrial
lipid CL in
PITP–/–
liver is consistent with energetic compromise
(Fig. 7C). The mass
increase of CL in
PITP–/–
brain may reflect compensatory attempts by this tissue to enhance
mitochondrial activity in response to an unfavorable energy status
(Fig. 7C).
|
Hypoglycemia in
PITP–/–
Mice—The aponecrotic cell death observed in the
PITP–/–
spinocerebellar system and liver diagnoses a systemic problem. In this regard,
we find
PITP–/–
neonates are severely hypoglycemic. We measure 9-fold reductions in serum
glucose levels of
PITP–/–
mice (9.6 ± 3.6 mg/dl) relative to
PITP+/+ siblings
(88.0 ± 6.6 mg/dl serum; Fig.
9A). This reduction in serum glucose is of similar
magnitude to that measured for
PITP+/+ mice that
are fasted overnight, despite the fact that stomachs of all
PITP–/–
mice analyzed were filled with milk at the time of analysis (not shown).
|
) mice with stomachs full of milk were fasted as indicated, sacrificed,
and liver glycogen measured. Mean glycogen values are given for
PITP
Reduction in serum glucose is not secondary to hyperinsulinemia.
Circulating insulin levels in
PITP–/–
mice are 9-fold lower than those of
PITP+/+ siblings
(0.5 ± 0.2 versus 4.5 ± 0.2 ng/ml, respectively;
Fig. 9A). Given the
hypoglycemia of
PITP–/–
mice, these low insulin levels are appropriate (see below). We also measured
an increase in circulating -hydroxybutyrate in
PITP–/–
versus PITP+/+ serum
(19.1 ± 1.2 and 11.2 ± 1.8 mg/dl, respectively;
Fig. 9A), and this
increase is recapitulated in serum of
PITP+/+ mice that
were fasted overnight (20.2 ± 0.1 mg/dl). -Hydroxybutyrate is
generated by mitochondrial oxidation of fatty acids and serves as an
alternative energy source for hypoglycemic tissue. Modest elevations are
scored for bilirubin and alkaline phosphatase in
PITP–/–
mice (consistent with modest hepatic stress), and a 2-fold increase in the
gluconeogenic enzyme aspartate aminotransferase is also measured
(Table I).
Reduced Proglucagon Gene Expression in
PITP–/–
Mice— Mice normally effect dramatic changes in circulating
insulin, corticosteroid, and glucagon levels to activate hepatic
gluconeogenesis late in gestation. This reprogramming of hormonal signaling
serves as a protective mechanism for the impending loss of the glucose source
provided by the maternal circulatory system in utero
(33). Mutant animals are
appropriately responding to hypoglycemia as evidenced by elevated levels of
circulating corticosteroids, a hormonal stimulus for gluconeogenesis. Levels
of plasma corticosteroid are highly elevated in
PITP–/–
mice relative to
PITP+/+ littermates
(51.64 versus 19.58 ng/ml, respectively). We also compared expression
of the gluconeogenic enzymes Glc-6-Pase and phosphoenolpyruvate carboxykinase
in PITP+/+ and
PITP–/–
mice. Expression of both genes in PITP
–/– liver compares
favorably to PITP+/+
control as their corresponding mRNA levels in
PITP+/+ liver are
1.3 ± 0.1- and 1.1 ± 0.2-fold increased relative to those of
PITP–/–
liver (Fig. 9B). Thus,
transcriptional control of key gluconeogenic enzymes also appears intact in
the mutant mice. Elevated serum levels of the gluconeogenic enzyme aspartate
aminotransferase in
PITP–/–
further support this conclusion (Table
I).
By contrast, proglucagon gene expression is inappropriately depressed in
PITP–/–
animals (Fig. 9B). By
using -actin mRNA as a normalizing factor, proglucagon gene expression
in pancreas is reduced 11.7 ± 0.1-fold in
PITP–/–
mice relative to
PITP+/+ siblings. In
independent agreement with the Northern data, we record a 5-fold reduction in
circulating glucagon levels in mutant animals (246.68 and 49.32 pg of
glucagon/ml of plasma for
PITP+/+ and
PITP–/–
mice, respectively).
Glycogen Metabolism in
PITP–/–
Mice—Normally, liver glycogen is rapidly consumed in response
to hypoglycemia. Reductions in plasma glucagon therefore predict defects in
glycogenolysis in
PITP–/–
mice. Periodate acid-Schiff staining and visualization of glycogen rosettes by
electron microscopy indicate substantial and inappropriate glycogen stores in
PITP–/–
liver (not shown). Quantitative analyses support these observations. As
expected, PITPa+/+ liver
contains 109.0 ± 15.3 µmol of glycogen-derived glucose/g of tissue,
and liver glycogen stores are depleted by a 10-h fast
(Fig. 9C). Strikingly,
the hypoglycemia of the mutant mice notwithstanding,
PITPa–/– liver
contains 107.0 ± 12.2 µmol of glycogen-derived glucose/g of tissue
(Fig. 9C). This
compares favorably to wild-type values.
To characterize further the derangement in glycogen metabolism, we tested
whether glycogen stores in
PITP–/–
liver are resistant to depletion by fast. We found that a 10-h fast evokes
complete depletion of glycogen from mutant liver
(Fig. 9C). When levels
of liver glycogen are measured as a function of duration of fast in
PITP–/–
and PITP+/+ animals,
we found the rate of glycogen depletion from fasted
PITP–/–
liver is delayed. Whereas
PITP+/+ mice deplete
50 and 70% of their liver glycogen stores by 2 and 4 h of fast,
respectively,
PITP–/–
mice effect only a modest 23 and 27% depletion in liver glycogen stores after
2 and 4 h of fast, respectively (Fig.
9D). Taken together, these results are not consistent
with an absolute block in
PITP–/–
liver glycogen catabolism. Rather, the data indicate
PITP–/–
liver exhibits kinetic defects in the rate of glycogenolysis and additionally
suggest an inappropriate channeling of glucose to glycogen synthesis in the
hypoglycemic mutant mouse.
Morphological Alterations in
PITP–/–
Pancreas—Reduced proglucagon gene expression could reflect
specific derangement of proglucagon gene expression in the mutant animals or
could arise indirectly from generalized pancreatic dysfunction. Although
exocrine pancreas morphologies of
PITP+/+ and
PITP–/–
mice are similar,
PITP–/–
pancreas exhibits obvious structural defects in that pancreatic islets are not
at all prominent. Serial sectioning of
PITP+/+ and
PITP–/–
pancreas, and quantification of total islets in the organ, revealed a greater
than 2-fold reduction in absolute islet number in
PITP–/–
pancreas relative to wild type (Fig.
10A). In addition to the significant reduction in the
number of islets, the morphology of identifiable islets in
PITP–/–
pancreas is also highly abnormal. Islet cells are sparse and shrunken, and
large vacuolations are abundant in the islets themselves
(Fig. 10B). The
damage of pancreatic islets in
PITP–/–
mice provides a direct rationale for reduced proglucagon expression and
defective gluconeogenesis in
PITP–/–
animals. Moreover, the very low levels of circulating insulin measured for
PITP–/–
mice (see above) may also result from islet -cell insufficiencies and
not simply from the hypoglycemic state of these animals.
|
PITP–/–
Cells Are Not Generally Defective in UPR—A linkage between
activation of UPR and survival of pancreatic secretory islet cells has been
demonstrated (34,
35). The ER abnormalities in
PITP-deficient enterocytes, hepatocytes, and neurons, when coupled with
loss of functional pancreatic islets in mutant mice, suggested the possibility
that
PITP–/–
cells are intrinsically defective in UPR.
First, we compared the response of
PITP+/+ and
PITP–/–
ES cells and MEFs to glucose deprivation. Cells defective in UPR are sensitive
to such a challenge (34). In
neither case do we record an unusual sensitivity of
PITP–/–
cells to glucose deprivation.
PITP+/+ ES cells
maintain levels of viability that are 71 ± 5 and 55 ± 8% of
control after 42 and 72 h of glucose starvation, respectively. The
corresponding viability levels for isogenic
PITP–/–
ES cells are 71 ± 4 and 54 ± 10%
(Fig. 10C).
Similarly, PITP+/+
MEFs exhibit viabilities of 82 ± 5 and 73 ± 10% after glucose
deprivations of 42 and 72 h, respectively. Isogenic
PITP–/–
MEFs exhibit corresponding viabilities of 84 ± 4 and 76 ± 8%
(Fig. 10C).
Second, 35S-labeled amino acid pulse-radiolabeling experiments
demonstrate that treatment of
PITP+/+ and isogenic
PITP–/–
ES cells with agents that activate UPR (i.e. dithiothreitol) evoke
reduced incorporation of 35S-labeled amino acids into nascent
protein in both wild-type and mutant ES cells. The rate of protein synthesis
in the face of dithiothreitol is 7.2 ± 0.7 and 5.2 ± 0.7% of
unchallenged control for
PITP+/+ and
PITP–/–
ES cells, respectively (Fig.
10D). This effect is a result of a UPR-mediated
translational block (34).
These findings are qualitatively reproduced in other experiments that employed
thapsigargin challenge to induce UPR (Fig.
10D).
Finally, we assessed whether challenge of
PITP+/+ and
PITP–/–
MEFs and ES cells with the UPR-activating agent tunicamycin leads to a
differential response in expression of the ER stress-induced transcription
factor CHOP and the chaperone protein calnexin. Expression of these two
polypeptides is potently enhanced by induction of the UPR. Thus, CHOP and
calnexin serve as reliable reporters of the UPR status in cells. As shown in
Fig. 10E, CHOP is
normally expressed at very low levels in
PITP+/+ and
PITP–/–
MEFs, but challenge with tunicamycin evokes a robust induction of CHOP
expression in MEFs of either genotype. Similarly, calnexin levels increase in
both PITP+/+ and
PITP–/–
MEFs upon tunicamycin challenge (Fig.
10E). Congruent results are recorded with
PITP+/+ and
PITP–/–
ES cells (not shown). We conclude
PITP–/–
cells are not only competent for induction of UPR but that the nullizygous
cells are not subject to constitutive activation of the UPR.
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
Loss of PITP
activity in mice manifests itself in spinocerebellar
disease characterized by reactive gliosis of the cerebellum and brain stem and
WM and GM deficits in spinal cord. The spinal cord pathologies include
infiltration by inflammatory cells, aponecrosis of motor neuron cell bodies in
the ventral horn, and pervasive demyelination that is particularly apparent in
the dorsal columns. PITP is also required for proper absorption of
dietary fat and fat-soluble vitamins through duodenal enterocytes into the
circulation and for proper lipid homeostasis in the liver. Finally,
PITP–/–
mice are severely hypoglycemic. The hypoglycemia is associated with reduced
proglucagon gene expression, compromise of pancreatic islet integrity, and
defects in liver glycogen metabolism. These findings suggest unanticipated
roles for PITP in signaling pathways that interface with trafficking of
luminal lipid cargo from the ER and glucose homeostasis in mammals.
PITP and Neonatal Survival—The development
of
PITP–/–
mice to term supports our finding that PITP-deficient ES cells maintain
pluripotency and competence for membrane trafficking and growth factor
signaling (17). Rather, the
neonatal mortality of
PITP–/–
mice indicates a role for PITP in regulating physiological functions
that are critical in the context of the whole mouse. We conclude PITP
nullizygosity manifests itself primarily upon extinction (by birth) of the
nutritional resource provided by maternal circulation in utero, and
reliance of the neonate on assimilation of its own dietary intake.
PITP Deficiency and Steatosis—The most
energy-rich component of the neonatal diet is TG, and the evidence suggests
PITP–/–
mice are unable to effectively avail themselves of this nutrient source. In
mammals, long chain TGs of maternal milk are hydrolyzed to monoacylglycerols
and fatty acids. These compounds are transported across the apical enterocyte
plasma membrane and reassembled into TG cores in the smooth ER lumen. The TG
cores are then co-assembled with other lipids, fat-soluble vitamins, and
apolipoproteins into chylomicrons either in the ER or in the Golgi complex.
Subsequently, chylomicrons are transported across the enterocyte basolateral
plasma membrane via the secretory pathway
(37). Enterocytes and
hepatocytes share analogous pathways for assembly and secretion of
lipoproteins into the circulation. The suckling-dependent microvesicular
steatosis of PITP–/– intestine and liver
suggests that PITP plays a common role in both intestinal and hepatic
pathways for secretion of dietary lipids. A failure in transport early in the
secretory pathway is indicated as electron microscopy suggests neutral lipids
accumulate in smooth ER of PITP nullizygous enterocytes and in
hepatocytes.
The intestinal steatosis of
PITP–/–
mice exhibits similarities with human CRD, a family of autosomal recessive
disorders. Human CRD patients are characterized by pediatric failure to
thrive, accumulation of chylomicrons within villus-associated enterocytes of
the duodenum, reductions in post-prandial plasma TG and -tocopherol
levels, and spinocerebellar involvement
(18,
19,
32,
38). Similarly,
PITP–/–
mice gain body mass slowly and are deficient in body fat. Moreover,
villus-associated
PITP–/–
enterocytes accumulate intracellular lipid in a suckling-dependent manner.
Finally,
PITP–/–
mice exhibit large reductions in both post-prandial plasma TG and in brain
-tocopherol. These data suggest
PITP–/–
mice hydrolyze dietary TG in the lumen of the duodenum, transport the products
into the enterocyte, and properly reassemble dietary TG in the ER lumen but
are defective in transport of re-esterified TGs across the
PITP–/–
enterocyte to the basolateral plasma membrane. The microvesicular hepatic
steatosis we observe in
PITP–/–
mice is not a symptom associated with human CRD, however.
PITP Deficiency and Glucose
Homeostasis—PITP–/–
neonates are hypoglycemic, even though key gluconeogenic enzymes appear to be
expressed normally. However, gluconeogenesis is an energetically expensive
process as 4ATP, 2GTP, 2NADH, and 4H2O are consumed in conversion
of 2 pyruvate units to 1 glucose molecule
(39). Fatty acid oxidation is
a potent engine that drives production of the ATP, GTP, and NADH utilized in
glucose synthesis and is also involved in production of ketone bodies that
serve as energy sources under conditions of low glucose availability. We
suggest that the inability to effectively deliver lipid from enterocytes and
hepatocytes to peripheral tissues imposes a heavy metabolic burden on
PITP–/–
neonates by compromising fatty acid oxidation as a pathway for powering both
gluconeogenesis and ketone production. Reductions in
PITP–/–
liver ATP are consistent with this interpretation of the data.
In addition,
PITP–/–
animals exhibit kinetic defects in glycogenolysis, and these maintain
substantial liver glycogen in the face of hypoglycemia. These defects coincide
with dramatic reductions in proglucagon gene expression in
PITP–/–
pancreas. Presently, we do not believe that dysregulation of proglucagon
expression reflects a direct transcriptional involvement of a nuclear
PITP pool in regulating proglucagon promoter function. Rather, we
hypothesize that glucagon deficit in
PITP–/–
mice more likely represents an indirect manifestation of the structurally
compromised endocrine pancreas that characterizes such mutants. Yet, because
the glycogen stores of
PITP–/–
liver are consumed during a modest fast, we conclude that regulation of
glucose allocation into the glycogen synthesis pathway must also be deranged
in
PITP–/–
mice.
PITP Deficiency and Integrity of the Spinocerebellar
System—PITP–/–
neonates develop a spinocerebellar neurodegenerative disease characterized by
aponecrotic cell death. In this regard, we find a correlation between site of
neuronal injury and regional specificity of dramatic reduction in cellular
ATP. Neonatal cerebellum may be particularly sensitive to injury in
PITP–/–
mice because it attempts to sustain the extraordinary rates of cell
proliferation that accompany the postnatal development of this organ in the
face of a challenging energetic environment. Similarly, Purkinje cells and
motor neurons are very large and metabolically active cells that are sensitive
to a variety of insults. We suggest
PITP–/–
Purkinje cell and motor neuron defects also represent manifestations of energy
deficit in these cells, thereby raising the possibility that a significant
component of the spinocerebellar disorder arises from the intrinsic
hypoglycemia and/or the intestinal and hepatic steatosis of
PITP–/–
mice.
Cellular Mechanisms for PITP Function in the
Mouse?— The lines of evidence linking PITP to potentiation
of PIP synthesis suggest that the tissue defects of
PITP–/–
mice result from deranged PIP metabolism. A demonstration that this is indeed
the case remains elusive as such derangements are not apparent in bulk PtdIns,
PtdIns-3-phosphate, PtdIns-4-phosphate, or PtdIns-4,5-bisphosphate levels in
PITP–/–
ES cells (17) or MEFs (this
study, not shown). Our data indicate that PITP does not play a critical
role in regulating dynamics of cell-essential phospholipid pools in most cell
types.
With regard to dysregulation of cellular homeostasis in PITP
nullizygous mice, we find that derangement of ER function is a common theme.
PITPs are classically invoked to function as lipid carriers that ferry PtdIns
from its ER site of synthesis to signaling membranes (plasma membrane and
Golgi) so that dynamic PIP pools can be replenished. In its simplest form,
this model predicts that PITP deficiency will compromise signaling in
membranes distinct from the ER. Although it remains possible that disruptions
of PITP-mediated signaling in non-ER compartments manifest themselves
indirectly in an ER pathology, the consistent smooth ER abnormalities of
PITP–/–
spinal and cerebellar neurons, enterocytes, and hepatocytes suggest that the
ER may itself represent a primary target of PITP function. In this
regard, PITP-dependent egress of PtdIns from the cytoplasmic leaflet of
ER membranes may be required for ER function via unanticipated mechanisms.
Alternatively, PITP may play unforeseen ER signaling functions that do
not require PITP-mediated transfer of lipid between organelles.
With regard to ER functions for PITP, we entertained the idea that
coupling of induction of UPR to ER secretory load is PITP-dependent and
is compromised in
PITP–/–
cells. Although our finding that
PITP–/–
ES cells and MEFs are UPR-competent argues against this idea, we cannot
exclude the possibility that UPR defects in
PITP–/–
mice are manifested only in professional secretory tissues (e.g.
pancreatic islet cells, Purkinje cells, enterocytes, and hepatocytes) where
high levels of cargo processing in the ER must occur efficiently.
An alternative possibility is that PITP plays a direct and specific
role in membrane trafficking from the ER in professional secretory cells by
playing a direct role in facilitating membrane trafficking at the level of
budding of specific ER-derived transport vesicles. This proposed activity is
congruent with the role of PITPs in yeast membrane trafficking
(1,
3–5).
The accumulation of dietary neutral lipids in the lumen of the enterocyte and
hepatocyte ER is consistent with such a function. Although constitutive and
regulated secretory pathway function is normal in
PITP–/–
ES cells (17), and development
of
PITP–/–
mice to term is also inconsistent with wholesale defects in membrane
trafficking from the ER of nullizygous cells, we propose that the ER membrane
trafficking function of PITP exhibits a large degree of cargo
specificity.
Is there independent supporting evidence for such a proposal? It was
recently demonstrated that inactivation of the SARA2 isoform of the
Sar1p GTPase forms a genetic basis for human CRD
(40). Sar1p is required for
the budding of COPII vesicles from ER membranes and for the concentration of
cargo into such transport vesicles
(41). Our data, when coupled
with the demonstrations of a role for Sar1p in human CRD, suggest the
attractive hypothesis that PITP regulates the Sar1p GTPase cycle in ER
membranes. We propose that a dedicated PITP/Sar1p interface is required
to optimize assembly and packaging of unusual luminal cargoes (such as large
TG cores) into ER-derived transport vesicles. This model shares common
features with previous proposals positing that the Sec14p PITP regulates the
activity of specific ADP-ribosylation factor GTPase-activating proteins
required for biogenesis of secretory cargo-laden transport vesicles from
trans-Golgi membranes in yeast
(6,
42).
|
FOOTNOTES |
|---|
* This work was supported in part by National Institutes of Health Grant NS37723 (to V. A. B.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
¶ Recipient of a National Institutes of Health grant.
To whom correspondence should be addressed. Tel.: 919-962-9870; Fax:
919-966-1856; E-mail:
vytas{at}med.unc.edu.
1 The abbreviations used are: PITPs, phosphatidylinositol transfer proteins;
CHOP, CCAAT/enhancer-binding protein homology protein; CL, cardiolipin; CRD,
chylomicron retention disease; DAG, diacylglycerol; ER, endoplasmic reticulum;
ES cells, embryonic stem cells; FFA, free fatty acids; GFAP, glial fibrillary
acidic protein; GM, gray matter; Glc-6-Pase, glucose-6-phosphatase; MEFs,
murine embryonic fibroblasts; PIP, phosphoinositide; PtdCho,
phosphatidylcholine; PtdIns, phosphatidylinositol; TG, triglyceride; TUNEL,
terminal deoxynucleo-tidyltransferase-mediated dUTP nick-end-labeling; UPR,
unfolded protein response; WM, white matter; CE, cholesteryl ester.
|
ACKNOWLEDGMENTS |
|---|
We acknowledge Douglas Cyr, Andrew Morris, Patrick Brennwald, Peter Petrusz, John York, Tushar Patel, William Snider, Virginia Godfrey, Scott Traske, and Rosalind Coleman for helpful discussions. We are particularly grateful to Gail Grossman for expert assistance in histochemistry; Hal Mekeel for helpful advice on electron microscopy; Patricia Gallagher for expert assistance in
-tocopherol measurements (University of North Carolina
Clinical Nutrition Research Center supported by National Institutes of Health
Grant DK56350); David Brenner and Joe Grisham for expert advice on liver
histopathology; Lisa M. Bain for brain histology; and Heather Collins and
Bryan Wolf (University of Pennsylvania Diabetes Center) for glucagon and
corticosteroid measurements. Support for body composition assays were provided
by grants from the National Institutes of Health to the University of Alabama
at Birmingham Clinical Nutrition Research Center. |
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|---|
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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