![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
|
|
|
|
|
||
Department of Animal and Nutritional Sciences, University of New Hampshire, Durham, NH 03824-3590
2To whom correspondence should be addressed. E-mail: dbobilya{at}cisunix.unh.edu.
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
KEY WORDS: • blood–brain barrier • brain capillary endothelial cells • zinc deficiency • zinc homeostasis • zinc transport
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
). Zinc deficiency during brain development in experimental animals causes permanent malformations (2
), which is presumably also true in humans (3
). A deficiency of zinc in young or mature individuals can cause brain abnormalities affecting neuromotor and cognitive function (4
). Brain dysfunction due to zinc deprivation may be indirectly due to a generalized decrease in zinc-dependent processes, such as protein synthesis, DNA and RNA synthesis, or cell membrane stability (3
). Zinc also has more specific neurological roles; neuronal zinc modulates the activity of neuropeptides that are released from synaptic boutons during nerve transmission (5
). Zinc deficiency is linked to altered neurotransmitter activity (6
). Excess extracellular zinc concentrations found in brain neurons have also been linked to neurodegenerative brain diseases, including Alzheimer’s disease (7
).
The concentration of zinc in brain tissue usually is not affected by zinc deficiency (8
–10
), although changes have been observed (11
). Brain zinc was also not influenced by a plasma zinc concentration that was 10 times above normal after infusion of a zinc acetate solution (12
). Nevertheless, the brain can suffer from zinc malnutrition, as evidenced by the alterations in cognition and behavior described above. Therefore, it is likely that these behavioral changes result from depletion of a relatively small, yet labile, zinc pool that could influence the extracellular brain fluid. The importance of zinc in brain function has been demonstrated; however, mechanisms by which this narrow range of extracellular zinc is maintained in the brain remain to be elucidated.
Whole-body zinc homeostasis is predominantly regulated by adjusting the efficiency of zinc absorption from the small intestine and the amount of endogenous zinc excretions into the gastrointestinal tract (13
,14
). These homeostatic mechanisms usually succeed in preventing the consequences of severe zinc malnutrition. Internal homeostatic mechanisms may also exist that regulate distribution of zinc inside the body. Such a system would enable the body to prioritize delivery of zinc to a tissue of relatively high priority, such as the brain, at the expense of low priority tissues, such as muscle or bone. For nutrient exchange to occur between blood and a tissue, the nutrient must pass through the blood vessel wall. We hypothesized that this barrier might possess regulatory capabilities and that the brain would likely be a tissue that possesses such homeostatic regulation.
We developed an in vitro model of the blood–brain barrier (BBB)3
to investigate the ability of this tissue to maintain brain zinc homeostasis. This in vitro model permits us to exquisitely manipulate the environment and measure functional responses of the cells. A similar model has frequently been used to investigate transport of drugs (15
,16
) into the brain. We exposed the model to environments that were moderately deficient or excessive in zinc. Extreme malnutrition was intentionally avoided because this provokes comprehensive pathological consequences that would have complicated interpretation of the data. Zinc transport kinetics were our principal functional parameters. We found that the capillary endothelial cells of the brain are responsive to zinc status—they adjusted their capacity to transport zinc, presumably to maintain brain zinc homeostasis.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
Brain capillary endothelial cells (BCEC) were isolated from Yucatan miniature swine (Sus scrofa) using methods previously described (17
,18
). All procedures were approved by the University of New Hampshire (UNH) Animal Care and Use Committee. Briefly, young (3- to 4-mo old) Yucatan miniature swine (Sus scrofa) from the UNH Miniature Swine Research Facility were anesthetized and subsequently euthanized by exsanguination. Approximately 20 g of cerebral cortical tissue were removed and washed with collection medium consisting of 2% fetal bovine serum (FBS) in HEPES (20 µmol/L) modification of minimum essential medium (MEM) with Earl’s salts, L-glutamine, and bicarbonate; plus 5 mg amphotericin B and 50 mg gentamicin/L. The tissue was mechanically dispersed into 1–2 mm3 pieces using sterile scalpel blades and pipet aspiration. Brain tissue was then dispersed in an equal volume of 1.0 g collagenase/L (type IA, 270 IU) in collection medium for a 60-min incubation at 37°C.
Nonendothelial cell types were removed from brain capillary fragments through successive centrifugation in a 25% albumin gradient, followed by filtering through screens of various pore sizes. Isolated brain capillary fragments were suspended in primary growth medium and seeded into fibronectin-coated (2 µg/cm2) tissue culture flasks for growth at 37°C, with 5% CO2 and 95% humidity in a water-jacketed incubator. Primary growth medium was composed of 15% platelet-poor horse serum (HS) in MEM with bicarbonate, 5 mg amphotericin B and 50 mg gentamicin/L plus 100 mg of heparin and 50 mg hypothalamic extract/L. Hypothalamic extract is an endothelial growth supplement that was prepared according to the methods of Maciag et al. (19
) and Thornton et al. (20
). The primary growth medium was changed daily until d 3, when the cells were switched to secondary growth medium consisting of 2% FBS and 13% HS in MEM, plus 5 mg amphotericin B, 50 mg gentamicin, 100 mg heparin, and 50 mg hypothalamic extract/L. BCEC were 55–65% confluent by d 4, when they were subcultured into experimental units.
Cell characterization.
BCEC isolations were monitored daily for endothelial characteristics, including density-inhibited growth and recognizable endothelial cell morphology: slightly ellipsoid nucleus surrounded by a cigar-like plasma membrane (21
). BCEC tested positive for acetylated-LDL (Ac-LDL) uptake (22
) and Factor VIII-related antigen (23
), classic endothelial cell characterization assays. Electrical resistance of the BCEC grown on the Transwell membranes was recorded daily as a physiological measure of monolayer integrity and barrier function (24
).
Alteration of zinc status.
Primary cultures of BCEC were seeded into fibronectin-coated (2 µg/cm2) experimental units. Cells used in zinc uptake studies were seeded at 12,000 cells/cm2 in 35-mm diameter tissue culture-treated dishes (Corning Costar Laboratory Science, Park Ridge, IL). Zinc transport studies used BCEC seeded at 85,000 cells/cm2 onto 12-mm Transwell cell culture inserts with 0.4-µm pore size (3401; Corning Costar Laboratory Science). BCEC were grown to confluence in designated treatment media: control (normal zinc), low zinc, zinc back, or excess zinc (high). All treatment media included 100 mg heparin and 50 mg endothelial cell growth supplement per liter. Treatment media were applied to cells on d 1 and 3, with zinc transport and uptake being measured on d 5.
The control medium was composed of 2% FBS and 13% HS in MEM. Zinc concentration was analyzed to be 3 µmol/L. Low zinc medium was composed of 2% dialyzed FBS and 13% dialyzed HS in MEM. (Procedure for dialyzing the serum to remove endogenous zinc is described below.) Low zinc medium had a zinc concentration of 1.5 µmol/L, by analysis (described below). Our goal was to induce moderate zinc malnutrition in the cells. Zinc back medium consisted of 2% dialyzed FBS and 13% dialyzed HS in MEM plus zinc chloride (ZnCl2). The addition of ZnCl2 adjusted the zinc concentration back to that of the control medium, 
3 µmol/L, by analysis. The zinc back treatment enabled us to distinguish between cellular responses related to decreased zinc concentrations, compared with other responses that might be a result of using dialyzed serums. High zinc medium was composed of 2% FBS and 13% HS in MEM, with ZnCl2 added to achieve 50 µmol/L zinc, by analysis.
The induction of an in vitro zinc deficiency included the use of low zinc medium with 2% FBS and 13% HS that was dialyzed against EDTA to remove endogenous zinc. This was necessary because serum was the predominant source of zinc in our culturing media. Dialysis procedures were adopted from McClung and Bobilya (25
). Mineral analysis determined FBS and HS zinc concentrations of 25 µmol/L and 7 µmol/L, respectively, following dialysis procedures, compared with 50 µmol/L for FBS and 12 µmol/L for HS in undialyzed serum.
Zinc concentration.
Zinc concentrations were analyzed by flame atomic absorption spectrophotometry (Smith Hieftje 12; Jarrell Ash, Franklin, MA). Reference standards were prepared with ZnCl2 in 0.1 mol/L hydrochloric acid diluted in deionized water in the linear range of 1–25 µmol/L zinc.
Zinc uptake (tissue culture dishes).
The rate of zinc uptake was measured using the procedures of Bobilya et al. (26
). Briefly, cells were washed three times with 2 mL HEPES buffer (10 mmol HEPES, 140 mmol NaCl, 7 mmol KCl, and 5.6 mmol glucose/L) at 37°C. The medium used to measure the kinetics of zinc uptake in all treatments of all experiments was the control growth medium. Because the transport medium was the same for all treatments, any differences in the rates of zinc uptake were due to differences in the cells because of their culturing environments. Control medium (3 µmol Zn/L) labeled with 1.0 µCi/mL (3.7 x 104 MBq/L) of 65Zn (Amersham, Arlington Heights, IL) at 37°C was applied to cells at 1 mL. BCEC were incubated for 30 min on an orbital shaker at 60 rpm in a 37°C incubator. Incubation medium was removed and cells were washed briefly (4–6 s) with 2 mL of HEPES/EDTA buffer (10 mmol EDTA, 10 mmol HEPES, and 150 mmol NaCl/L) at 4°C, followed by three washes with 2 mL of HEPES buffer at 4°C. Cells were solubilized for 60 min in 1.0 mL of 0.01% sodium dodecyl sulfate in 0.2 mol/L sodium hydroxide. Radioactivity was measured and the results converted to pmol Zn based on the specific activity of the labeled uptake medium. Samples were analyzed for protein concentration using the BCA method (27
). The rate of zinc uptake was expressed as pmol/(h x mg cellular protein).
Zinc uptake and transport (Transwell membranes).
Zinc uptake and transport across the BCEC were measured using methods adopted from Bobilya et al. (18
). Briefly, luminal and abluminal growth media were removed. BCEC monolayers on Transwell inserts were rinsed four times in 37°C HEPES buffer, followed by the application of 0.5 mL of 1.0 µCi/mL (3.7 x 104 MBq/L) 65Zn-labeled control medium (3 µmol Zn/L) at 37°C to the luminal chamber, and 1.5 mL of unlabeled control medium in the abluminal chamber. Cells were incubated for 60 min on an orbital shaker at 30 rpm inside a 37°C incubator. Abluminal medium was collected for analysis of radioactivity, and the results converted to pmol Zn based on the specific activity of the labeled transport medium. Zinc transport across the BBB was expressed as pmol Zn/(h x cm2 of cell monolayer). Zinc uptake and retention by the BCEC during the 60-min incubation was estimated by subsequently rinsing the Transwell insert twice in HEPES/EDTA buffer and then rinsing six times in HEPES buffer and removing the membrane with cells for analysis of radioactivity. The uptake results are expressed as pmol Zn/(h x cm2 of cell monolayer).
Statistical analysis.
Studies were analyzed as a randomized complete block, with each experiment as a block, with replicates over time as the block (28
). ANOVA and nonlinear fitting of the experimental data were performed with Systat (Version 9; Chicago, IL). Data from replications of the same experimental design were pooled when they passed the test of homogeneity. Fishers Protected Least Significant Difference test was used for pair-wise comparisons of multiple groups. Dunnett’s test was used for comparison of multiple groups with a control group. Differences were considered significant when P < 0.05.
Reagents.
Unless otherwise stated in the text, all reagents were obtained though Sigma (St. Louis, MO).
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
) in an effort to maintain cellular zinc homeostasis. This augmentation of zinc uptake occurs before a significant change in total zinc content can be measured. In our laboratory, the most sensitive indicator of cellular zinc status was the rate of zinc uptake.
Zinc uptake by the BCEC and transport from the luminal chamber (analogous to the blood) to the abluminal chamber (analogous to the brain) was measured during different time increments. The amount of zinc taken up and retained by the BCEC over time is presented in Figure 1
. The data represent the quantity of zinc taken up by the cells and retained, as determined by the 65Zn content of the cells after the incubation period. The kinetic data reflect two simultaneous phenomena and are best fit (R2 = 0.996) by a nonlinear equation that combines a saturation component with a linear component. We believe that the predominance of the saturation component during the initial 30 min reflects the zinc that rapidly enters the BCEC from the luminal side and moves through the cells to exit on the abluminal side; the result is an apparent saturation in the amount of zinc (65Zn) in the cells as the same amount exits on the abluminal side as enters from the luminal side. (It is also possible that some 65Zn exits back to the luminal chamber.) This equilibrium is achieved in 20 min, indicating that the rate at this time is the average speed at which zinc crosses the cell monolayer. The second component of zinc uptake and retention by the BCEC is linear and is clearly recognizable after 90 min; it reflects the zinc that enters the BCEC from the luminal side that is retained by the cells for their own purposes.
|
. Substantial amounts of zinc (65Zn) begin to be detected in the abluminal chamber after 20 min, and the amount increased linearly for at least 2 h. This reflects a constant rate of zinc delivery from the BCEC into the abluminal chamber, with no significant movement of 65Zn back into the cells or the luminal chamber. These results indicate that the optimal duration for measuring zinc transport across the BBB model would be 60–120 min, while the optimal incubation time for measuring the rate of zinc uptake would be 20–60 min. Therefore, the 20-min incubation time was used when we were only measuring zinc uptake (Fig. 7)
, but the 60-min incubation was used when we were simultaneously measuring zinc uptake into and transport across the cells (Figs. 3
4
5
6
).
|
|
|
|
|
|
. Zinc uptake did not differ between the two control groups (control and Zn back), demonstrating that the procedures for reducing the zinc concentration of the low Zn treatment medium did not introduce an artifact into the results. Zinc uptake was increased 9% (P < 0.05) by the low zinc environment relative to both control groups. The increase in zinc uptake under these moderately deficient conditions represents a significant response by the cells to their environmental zinc status. Because the same medium was used to measure zinc transport kinetics in all cells, regardless of the treatment, any difference in the transport rate between treatments reflects a difference in the cells’ capacity to transport zinc under the influence of the different zinc environments.
The moderate zinc deficiency also influenced zinc transport across the monolayer of BCEC grown on Transwell membranes in a BBB model (Fig. 4
). The capacity to transport zinc did not differ between the two control groups. The moderately low Zn treatment increased (P < 0.05) the rate of zinc transport 16% relative to both control groups. The physical integrity of the BBB model was not affected by the moderate zinc deficiency as determined by measurement of the barrier’s electrical resistance, which reflects its permeability to small ions.
The effects of moderate zinc excess on zinc transport across the BCEC grown on Transwell membranes are presented in Figure 5
. The rate of zinc transport across the BBB model was decreased 11% (P < 0.02) by growth in the high zinc environment. There were no changes in the physical integrity of the BBB model due to the high zinc environment, as determined by measurement of the barrier’s electrical resistance.
The moderately excessive zinc environment increased by 28% (P < 0.001) zinc uptake and retention by the BCEC on the Transwell membranes (Fig. 6
). Greater uptake in a higher zinc environment was counterintuitive, so we tested this finding using another model of the BCEC—this time growing the cells in culturing dishes rather than on Transwell membranes and measuring zinc uptake for 20 min (Fig. 7
). Once again, the high zinc environment increased (P < 0.001) the rate of zinc uptake 30%, confirming this remarkable observation.
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
) with our in vitro model of the BBB (17
) to test this hypothesis. Zinc transport by BCEC was affected by changes in their zinc status. The BCEC increased their capacity to acquire zinc in response to zinc restriction, but they also increased their release of zinc to the brain. To our knowledge, this is the first demonstration that blood vessel walls adapt to regulate tissue zinc homeostasis. The implication is that the endothelium is an important participant in the distribution of zinc among the tissues of the body.
One of our challenges was the establishment of a cell culture model system that would adequately reflect the BBB. Isolating BCEC and cultivating them into an in vitro BBB model has been a persistent challenge for the scientific community (15
,29
). We have been successful in isolating a very pure population of capillary fragments from pig brains. In culture, endothelial cells grew out from the capillaries and proliferated. We subcultured these cells into our model systems, so the cells used in these experiments were in passage 1 and 10 generations from their in situ progenitors. We estimated the purity of our cell populations to be 98–99% endothelial at the time of experimentation, with most of the nonendothelial cells being astrocytes or glial cells.
The quantity of new zinc entering into the endothelial cells of the BBB from the blood reached an initial plateau in 20 min. This plateau marks the point when the quantity of zinc entering the cells is roughly equivalent to the amount of zinc exiting the cells and indicates the amount of time required for zinc to pass though the BBB. This 20-min period is in close agreement with the time interval reported by other investigators for nutrient transport across an endothelial barrier (30
,31
). By 60 min, it is evident that a substantial portion of the zinc entering the cells remains within rather than being transported out to the other side of the barrier, presumably reflecting the portion of new zinc that is retained by the cell for metabolic purposes. Very little is known about the specific regulators of intracellular zinc trafficking (32
), although zinc status and physiological function would likely be factors.
Another of our challenges was the imposition of a moderate zinc deficiency on our model system. We considered it imperative for our model to retain physiological integrity to provide relevant data. Severe zinc deficiency disrupts BBB integrity in our model (data not shown), as well as an in vitro pulmonary endothelium (33
) and a rat brain model (11
). The numerous metabolic and physiologic pathways that are disrupted during severe zinc deficiency would diminish our ability to interpret the zinc transport data (34
). Therefore, we developed procedures to impose a moderate zinc deficiency on our BBB model. Our previous work with another cell line demonstrated that one of the early responses to zinc deficiency is an increase in the cell’s capacity to acquire zinc (25
). After much iteration, we established procedures that would impose an environment where the BCEC were zinc-deficient (demonstrated by an increase in their capacity to acquire zinc) while remaining healthy enough to perform important physiological functions (able to maintain BBB integrity). This demonstrated that the barrier formed by the brain endothelium can resist physical disruption by moderate zinc depletion.
The BCEC responded to our low zinc environment by enhancing their ability to acquire zinc from the blood. The 9% increase in uptake capacity of the BCEC is less than the 66% increase observed in zinc-deficient arterial endothelial cells (25
) or the 40-fold increase achieved in microbes (35
), but the BCEC in our model are not rapidly dividing (as the microbes) and are physiologically responsible for a tissue (the brain) with a relatively low rate of zinc turnover. Therefore, this adaptation reflects a significant improvement that would enable the cells to better meet their needs for zinc from the low zinc environment. Our procedures tend to understate the treatment differences because: 1) zinc uptake is partially confounded with zinc retention when measured in a 60-min incubation, and 2) the zinc concentration that would detect the greatest quantitative differences in transport rate would be at its Vmax (15–20 µmol/L) (18
), rather than at the much lower normal zinc concentration that was used in this study. In hindsight, this would have produced more impressive quantitative differences. This should not, however, diminish the importance of the present conclusions.
These cells of the BBB also enhanced their ability to transport zinc all the way across the cell monolayer, from the blood into the brain interstitium. Thus, the BCEC increased their export of zinc (into the brain) during a period when they were experiencing zinc deficiency. They were altruistically working to maintain brain zinc homeostasis. The capacity of the BBB to sustain brain zinc homeostasis in the face of persistent zinc deficiency is limited, as demonstrated by the many attendant neuropathologies (1
,36
). However, these adaptations by the BCEC would reduce the severity of these maladies.
The BCEC responded to our high zinc environment by reducing their transport of zinc across the cell monolayer, analogous to transport from the blood into the brain interstitium. This protective response would reduce the potentially toxic entrance of excessive amounts of zinc into the brain. Excess brain zinc has been implicated as a factor in ß-amyloid accumulation in Alzheimer’s disease (7
,37
).
The high zinc environment unexpectedly consistently and significantly increased the capacity for zinc uptake into the cells even though transport across the cells was reduced. This may be the cells’ effort to withdraw zinc from the environment to protect other cells of the brain. Alternatively, this may reflect an enhanced capacity of the cells to sequester or export zinc, serving to draw zinc in at a faster rate when subsequently exposed to the nonexcessive zinc concentrations we used to measure zinc uptake.
We are currently investigating the molecular mechanisms underlying these changes. Likely candidates would include a change in the quantity of the cell’s zinc import proteins (25
,35
,38
), zinc export proteins (13
,39
), zinc storage proteins (40
), or intracellular vesicles (41
). Metallothioneins appear to be important in maintaining essential zinc balance within neurological cells (42
). The importance of ZnT-3 zinc transporter and vesicular zinc remains elusive (43
). Elucidating these phenomena will increase our understanding of how zinc homeostasis is accomplished in cells and tissues, like the brain, and improve our ability to interpret evidence that results when zinc homeostasis is unsuccessful, i.e., during the neuropathologies characteristic of zinc deficiency and zinc toxicity.
|
ACKNOWLEDGMENTS |
|---|
|
FOOTNOTES |
|---|
3 Abbreviations used: BBB, blood–brain barrier; BCEC, brain capillary endothelial cell; FBS, fetal bovine serum; HS, horse serum; MEM, minimum essential medium; UNH, University of New Hampshire.
Manuscript received 11 April 2002. Initial review completed 1 May 2002. Revision accepted 24 May 2002.
|
LITERATURE CITED |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
1. Sandstead, H. H., Frederickson, C. J. & Penland, J. G. (2000) History of zinc as related to brain function. J Nutr. 130:496S-502S.
2. Dreosti, I. E., Manuel, S. J., Buckley, R. A., Fraser, F. J. & Record, I. R. (1981) The effect of late prenatal and/or early postnatal zinc deficiency on the development and biochemical aspects of the cerebellum and hippocampus in rats. Life Sci. 28:2133-2141.[Medline]
3. Keen, C. L., Taubeneck, M. W., Daston, G. P., Rogers, J. M. & Gershwin, M. E. (1993) Primary and secondary zinc deficiency as factors underlying abnormal CNS development. Ann. N. Y. Acad. Sci. 678:37-47.[Medline]
4. Henkin, R. I., Patten, B. M., Re, P. K. & Bronzert, D. A. (1975) A syndrome of acute zinc loss. Arch. Neurol. 32:745-751.
5. Frederickson, C., Suh, S., Silva, D., Frederickson, C. & Thompson, R. (2000) Importance of zinc in the central nervous system: the zinc-containing neuron. J. Nutr. 130:1471S-1483S.
6. Huntington, C., Shay, N., Grouzmann, E., Arseneau, L. & Beverly, J. (2001) Zinc status affects neurotransmitter activity in the paraventricular nucleus of rats. J. Nutr. 132:270-275.
7. Bush, A. I., Pettingell, W. H., Multhaup, G., Paradis, M. D., Vonsattel, J. P., Gusella, J. F., Beyreuther, K., Masters, C. L. & Tanzi, R. E. (1994) Rapid induction of Alzheimer AB amyloid formation by zinc. Science 265:1464-1467.
8. Wallwork, J. C., Milne, D. B., Sims, R. L. & Sandstead, H. H. (1983) Severe zinc deficiency: effects on the distribution of nine elements (potassium, phosphorus, sodium, magnesium, calcium, iron, zinc, copper and manganese) in regions of the rat brain. J. Nutr. 113:1895-1905.
9. Keen, C. J., Reinstein, N. H., Goudey- Lefevre, J., Lefevre, M., Lonnerdal, B., Schneeman, B. O. & Hurley, L. S. (1985) Effect of dietary copper and zinc levels on tissue copper, zinc, and iron in male rats. Biol. Trace Elem. Res. 8:123-136.
10. O’Dell, B. L., Becker, J. K., Emery, M. P. & Browning, J. D. (1989) Production and reversal of the neuromuscular pathology and related signs of zinc deficiency in guinea pigs. J. Nutr. 119:196-201.
11. Noseworthy, M. D. & Bray, T. M. (2000) Zinc deficiency exacerbates loss in blood-brain barrier integrity induced by hyperoxia measured by dynamic MRI. Proc. Soc. Exp. Biol. Med. 223:175-182.
12. Blair-West, J. R., Denton, D. A., Gibson, A. P. & McKinley, M. J. (1990) Opening the blood-brain barrier to zinc. Brain Res. 507:6-10.[Medline]
13. McMahon, R. J. & Cousins, R. J. (1998) Regulation of the zinc transporter ZnT-1 by dietary zinc. Proc. Natl. Acad. Sci. USA 95:4841-4846.
14. King, J., Shames, D. & Woodhouse, L. (2000) Zinc homeostasis in humans. J. Nutr. 130:1360S-1366S.
15. Cecchelli, R., Dehouck, B., Descamps, L., Fenart, L., Buee-Scherrer, V., Duhem, C., Lundquist, S., Rentfel, M., Torpier, G. & Dehouck, M. P. (1999) In vitro model of evaluating drug transport across the blood-brain barrier. Adv. Drug Deliv. Rev. 36:165-178.[Medline]
16. Janigro, D., Leaman, S. M. & Stanness, K. A. (1999) Dynamic in vitro modeling of the blood-brain barrier: a novel tool for studies of drug delivery to the brain. Pharm. Sci. Technol. Today 2:7-12.[Medline]
17. Bobilya, D. J., D’Amour, K., Palmer, A., Skeffington, C., Therrien, N. & Tibaduiza, E. C. (1995) Isolation and cultivation of porcine brain capillary endothelial cells as an in vitro model of the blood-brain barrier. Methods Cell Sci. 17:25-32.
18. Bobilya, D. J., Guerin, J. L. & Rowe, D. J. (1997) Zinc transport across an in vitro blood-brain barrier model. J. Trace Elem. Exp. Med. 10:9-18.
19. Maciag, T., Cerundolo, J., Ilsley, S., Kelley, P. R. & Forand, R. (1979) An endothelial cell growth factor from bovine hypothalamus: identification and partial characterization. Proc. Natl. Acad. Sci. USA 76:5674-5678.
20. Thornton, S. C., Mueller, S. N. & Levine, E. M. (1983) Human endothelial cells: use of heparin in cloning and long-term serial cultivation. Science 222:623-625.
21. Haudenschild, C. C. (1984) Morphology of vascular endothelial cells in culture. Jaffe, E.A. eds. Biology of Endothelial Cells 1984:129-140 Martinus Nijhoff Boston, MA. .
22. Voyta, J. C., Via, D. P., Butterfield, C. E. & Zetter, B. R. (1984) Identification and isolation of endothelial cells based on their increased uptake of acetylated-low density lipoprotein. J. Cell Biol. 99:2034-2040.
23. Hoyer, L. W., De Los Santos, R. P. & Hoyer, J. R. (1973) Antihemophilic factor antigen: localization in endothelial cells by immunofluorescent microscopy. J. Clin. Invest. 52:2737-2744.
24. Sill, H. W., Butler, C., Hollis, T. M. & Tarbell, J. M. (1992) Albumin permeability and electrical resistance as means of assessing endothelial monolayer integrity in vitro. J. Tissue Cult. Methods 14:253-258.
25. McClung, J. P. & Bobilya, D. J. (1999) The influence of zinc status on the kinetics of zinc uptake into cultured endothelial cells. J. Nutr. Biochem. 10:484-489.[Medline]
26. Bobilya, D. J., Briske-Anderson, M. & Reeves, P. G. (1992) Zinc transport into endothelial cells is a facilitated process. J. Cell. Physiol. 151:1-7.[Medline]
27. Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Provenzano, M. D., Fujimoto, E. K., Goeke, N. M. & Klenk, D. C. (1985) Measurement of protein using bicinchoninic acid. Anal. Biochem. 150:76-85.[Medline]
28. Gill, J. L. (1978) Design and Analysis of Experiments in the Animal and Medical Sciences 1978 Iowa State University Press Ames, IA. .
29. Gaillard, P., Voorwinden, L., Nielsen, J., Ivanov, A., Atsumi, R., Engman, H., Ringbom, C., De Boer, A. & Breimer, D. (2001) Establishment and functional characterization of an in vitro model of the blood-brain barrier, comprising a co-culture of brain capillary endothelial cells and astrocytes. Eur. J. Pharm. Sci. 12:215-222.[Medline]
30. Ghitescu, L. & Bendayan, M. (1992) Transendothelial transport of serum albumin: a quantitative immunocytochemical study. J. Cell Biol. 117:745-755.
31. Rowe, D. J. & Bobilya, D. J. (2000) Albumin facilitates zinc acquisition by endothelial cells. Proc. Soc. Exp. Biol. Med. 224:178-186.
32. Eide, D. J. (2000) Metal ion transport in eukaryotic microorganisms: insights from Saccharomyces cerevisiae. Adv. Microb. Physiol. 43:1-38.[Medline]
33. Hennig, B., Wang, Y., Ramasamy, S. & McClain, C. J. (1992) Zinc deficiency alters barrier function of cultured porcine endothelial cells. J. Nutr. 122:1242-1247.
34. MacDonald, R. S., Wollard-Biddle, L. C., Browning, J. D., Thornton, W. H., Jr. & O’Dell, B. L. (1998) Zinc deprivation of murine 3T3 cells by use of diethylenetrinitrilopentaacetate impairs DNA synthesis upon stimulation with insulin-like growth factor-I (IGF-I). J. Nutr. 128:1600-1605.
35. Zhao, H. & Eide, D. (1996) The yeast ZRT1 gene encodes the zinc transporter protein of a high-affinity uptake system by zinc induction. Proc. Natl. Acad. Sci. USA 93:2454-2458.
36. Golub, M. S., Takeuchi, P. T., Keen, C. L., Gershwin, M. E., Hendrickx, A. G. & Lonnerdal, B. (1994) Modulation of behavioral performance of prepubertal monkeys by moderate dietary zinc deprivation. Am. J. Clin. Nutr. 60:238-243.
37. Cherny, R. A., Atwook, C. S., Xilinas, M. E., Gray, D. N., Jones, W. D., McLean, C. A., Barnham, K. J., Volitakis, I., Fraser, F. W., Kim, Y. S., Huang, X., Goldstein, L. E., Moir, R. D., Lim, J. T., Beyreuther, K., Zheng, H., Tanzi, R. E., Masters, C. L. & Bush, A. I. (2001) Treatment with a copper-zinc chelator markedly and rapidly inhibits ß-amyloid accumulation in Alzheimer’s disease transgenic mice. Neuron 30:665-676.[Medline]
38. Grotz, N., Fox, T., Connolly, E., Park, W., Guerinot, M. L. & Eide, D. (1998) Identification of a family of zinc transporter genes from Arabodopsis that respond to zinc deficiency. Proc. Natl. Acad. Sci. USA 95:7220-7224.
39. Palmiter, R. D., Cole, T. B. & Findley, S. D. (1996) ZnT-2, a mammalian protein that confers resistance to zinc by facilitating vesicular sequestration. EMBO J 15:1784-1791.[Medline]
40. Coyle, P., Philcox, J. C. & Rofe, A. M. (2000) Zn-depleted mice absorb more of an intragastric Zn solution by a metallothionein-enhanced process than do Zn-replete mice. J. Nutr. 130:835-842.
41. Wenzel, H. J., Cole, T. B., Born, D. E., Schwartzkroin, P. A. & Palmiter, R. D. (1997) Ultrastructural localization of zinc transporter-3 (ZnT-3) to synaptic vesicle membranes within mossy fiber boutons in the hippocampus of mouse and monkey. Proc. Natl. Acad. Sci. USA 94:12676-12681.
42. Hidalgo, J., Castellano, B. & Campbell, I. L. (1997) Regulation of brain metallothioneins. Curr. Top. Neurochem. 1:1-26.
43. Cole, T., Martyanova, A. & Palmiter, R. (2001) Removing zinc from synaptic vesicles does not impair spatial learning, memory, or sensorimotor functions in the mouse. Brain Res. 891:253-265.[Medline]
This article has been cited by other articles:
|
|
 |
|
  W. Chowanadisai, S. L. Kelleher, and B. Lonnerdal Zinc Deficiency Is Associated with Increased Brain Zinc Import and LIV-1 Expression and Decreased ZnT-1 Expression in Neonatal Rats J. Nutr., May 1, 2005; 135(5): 1002 - 1007. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
