Differentialion (1992) 51 : 195-200

Differentiation Ontogeny, Neoplasia and Differentiation Therapy

0 Springer-Verlag1992

Time dependency of 1,25(OH)2D3induction of calbindin mRNA and calbindin expression in chick enterocytes during their differentiation along the crypt-villus axis Julie C.Y. Wu, Michael W. Smith, and David E.M. Lawson Department of Cell Biology, AFRC Institute of Animal Physiology and Genetics Research, Babraham, Cambridge CB2 4AT, UK Accepted in revised form July 7, 1992

Abstract. Quantitative methods of in situ hybridization and immunocytochemistry have been used to measure 1,25 dihydroxyvitamin D3 (1,25(OH),D3) induction of calbindin mRNA and calbindin protein expressed in jejunal enterocytes at all points along the crypt-villus axis over a 24 h period. Small amounts of calbindin mRNA detected in vitamin D, deficient (D-deficient) chick intestine increased rapidly to maximal values 8 h after hormone injection. The magnitude of this response was inversely related to age of enterocyte measured separately by injecting tritiated thymidine into D-deficient and 1,25(0H),D,-injected birds. Enterocytes of all ages expressed small amounts of calbindin 3 h after hormone injection. This amount of calbindin then increased up to 24 h after hormone injection. Maximal calbindin expression took place in basal villus enterocytes. Later decrease in the ability of upper villus enterocytes to express calbindin was associated with a similar fall in calbindin mRNA expression. Previously it was suggested that inefficient translation to calbindin mRNA might take place in basal villus enterocytes 48 h after vitamin D injection. Present work using 1,25 (OH),D, shows that calbindin expression takes place at a constant rate during this early stage of enterocyte development. Secondary events limiting higher rates of calbindin synthesis in upper crypt and basal villus enterocytes remain to be identified.

Introduction The steroid hormone 1,25-dihydroxyvitamin D, [1,25(OH),D,] has long been known to regulate calcium metabolism in obvious target tissues including bone, kidney and the intestine. Other work carried out recently shows this hormone to be responsible for the upregulation or inhibition of a large number of genes affecting Correspondmce t o : M.W. Smith

cell proliferation and development [ 101. Initial control over gene expression in this case involves interaction of the hormone with specific DNA binding proteins [6]. Mechanisms controlling gene transcription and translation have been studied mainly for the vitamin D induced protein, calbindin. Upregulation of calbindin expression correlates well with the maintenance of high levels of calcium absorption in the small intestine [I, 15, 161. Calbindin in this case is thought to facilitate calcium transport indirectly by acting as an intracellular buffer [13]. Timed studies of 1,25(OH),D, induction of calbindin expression in vitamin D deficient chick intestine show this molecule to appear first in lower villus and crypt enterocytes [22], an area also show by immunocytochemistry to contain the greatest amounts of vitamin D receptors [4]. Recent analysis also shows that this is an area where calbindin mRNA is expressed in highest amounts after vitamin D injection into vitamin D deficient chicks [8]. It is interesting to note that this is also the place where enterocytes complete differentiation of structure and digestive enzyme function during migration from the base of crypts to the tips of villi [17]. Taken together these findings suggest that 1,25(OH)2D3 might be involved in controlling the general pattern of cell differentiation taking place in this restricted population of enterocytes, in which case only part of this supposed action will be concerned with regulating calcium metabolism. New techniques of quantitative immunocytochemistry and in situ hybridization now make it possible to measure temporal changes taking place in gene expression during enterocyte development [8,19]. Present work uses this method of analysis to determine the time dependency of 1,25(OH),D3 induction of calbindin mRNA and calbindin expression in enterocytes migrating along chicken intestinal villi. Attempts are also made to distinguish the initial rates of calbindin expression from the long term accumulation of calbindin within enterocytes exposed to 1,25(OH),D3 before discussing possible mechanisms responsible for controlling calbindin synthesis.

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Methods Exper$mentu/. One-day-old Rhode Island Red x Light Sussex chicks maintaincd on a standard vitamin-D-deficient diet with free access to water for a period of 4 weeks [9] where either used as colitrols or killed for experiment 3, 8 or 24 h after the intracardiac injection of 125 ng 1,25(OH)zD3 dissolved in 0.1 mi of a 50: 1 mixture of propyleneglycol: ethanol to study the time course of calbindin mRNA and calbindin expression. A second group of vitamin D deficient chicks used as controls or given 4 daily injections of 125 ng 1,25(OH)2D, were killed for experiment 2, 24, 48, 72 and 96 h after being injected intramuscularly with tritiated thymidine (1 pCi/g body weight) to measure enterocyte migration rates. Pieces of proximal jejunal tissue removed from birds killed by cervical dislocation were in all cases flushed thoroughly with phosphate buffered saline to remove intestinal contents before being processed as described below. In situ hybridization. Tissue frozen in isopentane cooled in liquid N, was sectioned at 15 pni for later fixation in paraformaldehyde at room temperature. These sections were then washed in 0.2 M HC1 for 10 min to destroy endogenous alkaline phosphatase activity before being incubated for 15 min in a solution containing acetic anhydride and triethanolamine to reduce background staining. Later dehydration of these sections in alcohol and delipidation in chloroform was followed by a 10 min incubation in hybridization buffer containing an alkaline phosphatase-linked antisense oligonucleotide probe to detect calbindin mRNA. Reacted sections, washed repeatedly in hybridization and TRIS-containing buffers to remove excess probe, were finally incubated at room temperature for 16 h in a solution containing 5-bromo-4-chloro-3-indolyl phosphate and nitroblue tetrazolium to determine alkaline phosphatase activity in individual villus-attached enterocytes by microdensitometry. Controls were incubated under identical conditions using tissue taken from D-deficient chickens. The amount of mRNA in reacted sections was then determined by microdensitometry using a wavelength of 540 nm. Further details of this method have already been published [8]. Immunocytochemistry. The method used to determine calbindin involved initial fixation of tissue for 1 h at room temperature in a glutaraldehyde-paraformaldehyde containing buffer for subsequent embedding in Lowicryl K4M resin. Sections of this material, cut at lpm onto water, were dried at 60" C for initial 10 min incubation in medium containing thimerosal and bovine serum albumin to block non-specific binding of rabbit antichicken calbindin antibody to the tissue, berore being incubated with primary antibody to calbindin for 2 h. Washed sections incubated 30 inin with goldlabelled antirabbit IgG secondary antibody were later reacted with silver-enhancing reagents in order to carry out quantitative measurements of calbindin protein in villus-attached enterocytes by microdensitometry. Further details concerning the source of antibodies and the method uscd to carry out this procedure can be obtained from previous publications [8, 201.

1 ,25(OH),D3 injection were corrected for decay using the equation c = af - a, + k(af + ai/2) (tf - t,), where c is the correction to be applied to the final concentration; a, and ai are the final and initial concentrations respectively; ti and tf are the initial and final times when calbindin is measured; k is the apparent rate constant for calbindin degradation (0.024 h-'; t1,2 of 29 h [14]). In this case it is assumed that the rate of increase in calbindin concentration is constant between times 0-3, 3-8 and 8-24 h. The fact that it takes 2 h for calbindin to appear after 1,25(OH),D, injection [ l l ] does not materially affect negligible corrections applied over the initial 0-3 h period. Rates of calbindin synthesis calculated by regression analysis before and after correction give means & SEM. Other mean results are given & SEM. The'statistical significance of differences between mean values has also been assessed using paired and unpaired Student's t-tests. Muteriais. The 30 mer antisense alkaline phosphatase linked oligonucleotide probe used to measure calbindin mRNA came from Molecular Biosystems, (San Diego, USA). The artificial alkaline phosphatase substrate 5-bromo-4-chloro-3-indolyl phosphate and the nitroblue tetrazolium coupling reagent used to visualize this probe was purchased from Boehringer, (Mannheim, FRG). The secondary gold labelled antibody and silver enhancing kit used to visualize the primary antibody to chick calbindin and the [methyl-3H]thymidine (41 Ci/mmol) used to determine enterocyte migration rates both came from Amersham International, (Aylesbury, Bucks, UK). All other reagents used were of analytical grade.

Results

Intestinal structure and enterocyte kinetics Vitamin D injected into vitamin D deficient chicks has been shown previously to increase both the length of villi and the rate a t which enterocytes migrate along the crypt-villus axis [21]. Preliminary experiments were 0.4

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Thymidine lubelling experiments. Tissues used to determine enterocyte migration rates were fixed in sucrose-glutaraldehyde buffer for later embedding in glycolmethacrylate. Sections prepared from embedded tissue were then exposed to Ilford K 2 emulsion for up to 28 days before being developed in Ilford Phen-X developer. Silver grains over labelled cells were then enhanced by immersion for 1 min in silver intensifier solution (IN-5, Kodak). Tissue structure was at the same time made visible by staining lightly with eosin. Subsequent measurement of distances migrated by leading labelled enterocytes from their point of origin in the crypt and further measurements of crypt depth and villus height were carried out by image analysis (Magiscan 2A, Joyce-Loebl, Newcastle). This method is similar to that described previously [7]. Mathematical treutment of results. Calbindin concentrations recorded as arbitrary units of absorbance 0, 3, 8 and 24 h after

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Fig. 1. Enterocyte migration along chick jejunal villi. Vitamin D deficient chickens given 4 daily injections of 1,25(OH),D3 (0)or vehicle alone ( 0 )were killed at different times after thymidine injection to determine enterocyte migration rates. Each value gives the mean+ S.E.M. of the distance travelled by enterocytes measured at different times after thymidine injection ( 6 5 birds per point). The regression line gives a mean enterocyte migration rate of 3.82 & 0.22 pm/h

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Fig. 2. Photomicrographs showing distribution patterns of calbindin mRNA (a, c, e, g) and calbindin protein (b, d, f, h) in crypts and along villi of chicken small intestine. Chickens fed a vitamin D deficient diet (a, b) were killed 3 (c, d), 8 (e, f) and 24 h (g,

h) after the intramuscular injection of 125 ng 1,25(OH),D3. The

carried out to test the possibility that similar effects might occur when using 1,25(OH),D3 to induce calbindin synthesis. Vitamin D deficient chicks were either used as control or injected with 1,25(OH)2D, for a period of 4days before being killed for experiment. During this time tritiated thymidine was injected into both groups of birds to determine enterocyte migration rate. Image analysis of autoradiographs prepared from sectioned tissue showed 1,25(OH)2D, to have no significant effect on intestinal structure (villus heights of 776 & 82 and 857 f 93 pm; crypt depths of 93.0k4.2 and 99.7+ 3.9 pm for control and test birds respectively). The time course of enterocyte migration taking place under these conditions is shown in Fig. 1. Cells labelled in the crypts 2 h after injecting tritiated thymidine took about 20 h to reach

the crypt-villus junction. Migration along villi was linearly related to time up to 96 h after thymidine injection. Enterocyte migration rates calculated by regression analysis of results obtained from control and 1,25(OH)*D, injected birds were not significantly different (3.99 k0.34 and 3.64 0.26 pm/h respectively). The amount of vitamin D found previously to increase enterocyte migration rate was 5-25 times higher than the daily minimal requirement [21]. Inability of 1,Z(OH),D, to produce similar effects is probably related to the small amounts used in the present work. Analysis of pooled results gave a mean migration rate of 3.82 f0.22 pm/h. This value, which is assumed to apply along the whole length of the villus, was used subsequently to convert positional information obtained for calbindin synthesis to the age of enterocyte performing this function.

arrow shows the position for first detection of calbindin mRNA (c). Maximal expression of calbindin mRNA and calbindin are shown by white bars (e & h respectively). Scale bar (a); 100 pm

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Temporal aspects of 1, 2 5 ( 0 H ) 2D,-induced calbindin gene expression in chick jejunaE enterocytes A qualitative impression of the sequential way in which calbindin mRNA and calbindin protein becomes expressed at different positions along the crypt-villus axis during 24 h after 1,25(OH),D, injection is shown in Fig. 2. The apparent absence of calbindin mRNA and calbindin protein in tissue taken prior to l ,25(OH),D3 injection (Fig. 2 a & b respectively) confirms results published previously for tissue taken from vitamin D deficient chicks [8]. Small amounts of calbindin mRNA can however be detected in the lower part of villi 3 h after hormone injection (Fig. 2c). No apparent expression of calbindin can be detected under these conditions (Fig. 2d). Very large amounts of calbindin mRNA are present in enterocytes located all along the crypt-villus axis 8 h after hormone injection (Fig. 2e). Expression

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of calbindin is now also obvious particularly in the lower half of the villus (Fig. 2g). Calbindin detected 24 h after 1,25(OH),D3 injection is widely distributed along the whole length of the crypt-villus unit (Fig. 2h). Maximal calbindin levels in this case appear to be in mid-villus enterocytes. Expression of calbindin mRNA decreases in tissue taken from 24 h injected chickens (compare Fig. 2g and Fig. 2e). A quantitative analysis of these findings is illustrated in Fig. 3 . Trace amounts of calbindin mRNA detected in crypt and basal villus enterocytes in vitamin D deficient birds were not associated with any detectable expression of calbindin (Fig. 3 ; 0 hj. Small amounts of calbindin were however detected 3 h after injection of 1,25(OH)2D,. Calbindin mRNA levels at this time were highest around the crypt-villus junction. This pattern of mRNA expression is also seen 8 and 24 h after 1,25(OHj,D3 injection. Calbindin levels also increase at these later times with the greatest amounts of protein being eventually located in the mid villus region (Fig. 3 ; 24 hj. Lower levels of calbindin detected in upper villus enterocytes are closely associated with decreasing levels of calbindin mRNA suggesting that calbindin synthesis in this region might be under transcriptional control. A more detailed analysis of how 1,25(OHj,D3 injection affects calbindin synthesis can be carried out after transforming positional data onto a timescale. An example of how this operates for an enterocyte crossing the

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Fig. 3. Positional dependence of 1,25(OH),D, effects on calbindin mRNA and calbindin expression by chicken enterocytes. Tissues taken from vitamin D deficient chickens before and 3, 8 and 24 h after intracardiac injection of 125 ng 1,X(OH),D, were sectioned and processed as described in the text to determine calbindin (CaBP; 0 ) and calbindin mRNA levels (CaBP m R N A ; -). Values of optical absorbance in arbitrary units (a.u.) give means from analyses carried out on 6-24 villi (2-8 birds). Arrows show the position of the crypt-villus junction. Horizontal lines attached to the arrows show distances travelled by enterocytes 3, 8 and 24 h after 1,25(OH),D3 injection

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Fig. 4. Relating 1,25(OH),D, induced expression of calbindin mRNA to calbindin synthesis in individual chicken enterocytes. Results taken from Fig. 2 were used to calculate rates of calbindin synthesis (CaBP; A) as described in the text. Corresponding mean mRNA levels (CaBP m R N A ; o ) are also plotted for comparison. The UPYOM' shows the position of the crypt-villus junction. The ,figure inset shows increasing calbindin levels measured at different times (T) after 1,25(OH),D, injection before (0)and after ( 0 ) correction for calbindin degradation assuming a half-life time of 29 h

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crypt-villus junction at the time of 1,25(OH),D3 injection is shown in Fig. 3. Such an enterocyte will have migrated 8, 30 and 90 pm along the villus by the time the birds are killed 3,8 and 24 h later. Comparing calbindin levels at these different positions then allows one to estimate the time course for calbindin appearance over a period of 24 h. Results obtained from carrying out one such comparison is shown in Fig. 4. Initial rapid increase in calbindin levels continues at a slightly slower rate 8-24 h after 1,25(OH),D3 injection (Fig. 4; inset). Regression analysis of this data gives a rate of net synthesis of 9.5 5 1.4 a.u./h (a.u., arbitrary units; t=6.7; P=0.02). Calbindin is however known to be degraded with a half-life of 29 h in this tissue [14]. This value has therefore been used in the present work to correct original data for calbindin turnover (solid circles; Fig. 4 inset). Regression analysis of this data gives a corrected rate of calbindin synthesis of 12.9& 1.O a.u/h ( t = 12.5; P=O.O06). Similar estimates of calbindin synthesis carried out for enterocytes situated at different points along the villus at the time of 1,25(OH),D3 injection are plotted in Fig. 4. Enterocytes situated 100300 pm from the crypt-villus junction have the highest rates of calbindin production. The ability of cells to respond to 1,25(OH),D3 declines as enterocytes continue to migrate to the villus tip. Calbindin synthesis in enterocytes near the villus tip is however still half that found at the villus base. Mean mRNA levels responsible for calbindin synthesis can also be calculated for enterocytes located at different points along the villus. Results obtained from carrying out these calculations are compared with corresponding rates of calbindin synthesis in Fig. 4. The ability of enterocytes to express calbindin mRNA and synthesize calbindin show the same proportional decrease from mid to upper regions of the villus (170 to 128 a.u. and 11.4 to 8.9 a.u/h; 25 and 22% decrease respectively for enterocytes situated 300 and 700 pm from the cryptvillus junction). Similar comparisons made over the lower half’ of the villus show a marked discrepancy between the ability of enterocytes to express mRNA and synthesize calbindin (doubling of calbindin mRNA from 170 to 350 a.u. associated with a decrease in calbindin synthesis from 11.4 to 9.4 a.u/h for enterocytes situated 300 and 11 pm from the crypt-villus junction). These latter results suggest that a post-transcriptional control exists over calbindin synthesis in basal villus enterocytes. Discussion

Measuring the time course of 1,25(OH),D3 induction of calcium transport, calbindin mRNA expression and calbindin appearance in chick intestine has been used frequently in the past to investigate the part played by calbindin in enhancing calcium transport. Further studies have been carried out to determine whether 1,25(OH),D3 has any further action on calbindin synthesis beyond that associated with increased expression of the calbindin gene. The present view to emerge from this work is that calbindin acts more to maintain than

initiate increased calcium transport, possibly by acting as an intracellular carrier or secondary buffer protecting the cell from this potentially harmful cation [ 12, 13, 161. There is also some doubt as to whether 1,25(OH),D3 induced calbindin synthesis is controlled solely at the level of gene transcription [5,11,23]. All of these conclusions are however based upon work carried out on mucosal homogenates and this gives no positional information on possible changes taking place during enterocyte development. Recent work showed that mRNA/calbindin ratios were unusually high in basal villus enterocytes after injecting vitamin D [8] and this is confirmed in the present work when analyzing tissue taken 3-24 h after injecting 1,25(OH)2D3 into D-deficient chicks (Fig. 3). This ratio is however much reduced in 24 h compared with 3 or 8 h injected birds, mainly because of a continual increase in calbindin content. This is to be expected for any protein having a slow rate of turnover in migrating enterocytes. Absence of calbindin in vitamin D deficient chicks coupled with the ability of 1,25(OH),D3 to induce its expression without affecting enterocyte kinetics provides a unique opportunity to study the time dependency of events taking place in villus-attached enterocytes of different ages. Tracing the predicted history of cells during 24 h then allows one to construct profiles of calbindin appearance. Rates of calbindin appearance calculated from these profiles, further corrected for calbindin halflife, give more accurately determined rates of calbindin synthesis (Fig. 4). This correction is however more a matter of form than necessity since it applies equally well to enterocytes of all ages studied over the 24 h period. Results obtained show all enterocytes in the lower half of the villus to synthesize calbindin at about the same rate following hormone injection. Enterocytes in the upper half of the villus slowly lose this capacity to respond to hormone. A similar but less well defined ability of lower villus enterocytes to produce equal amounts of calbindin after 1,25(OH),D3 injection has been reported previously using cell fractions prepared from rat small intestine [ls]. Enterocyte levels of calbindin mRNA also vary 0-24 h after injecting 1,25(OH),D3 into D-deficient chicks. In this case it is assumed that calbindin synthesis is directly related to the mean level of calbindin mRNA measured over different time periods. The close agreement found between these two estimates supports the idea that calbindin synthesis is under transcriptional control in upper villus enterocytes (Fig. 4). This relationship breaks down in lower villus enterocytes, apparently because these cells are already synthesizing calbindin at near maximal rates. Post-transcriptional control over calbindin synthesis appears to operate in this particular region of the crypt-villus axis. Present results bear no relation to those obtained under steady state conditions where a similar profile for mRNA expression is associated with a continuing increase in calbindin level from bottom to top of villi in egg laying hens (Wu 8z Smith, unpublished findings). In this case villus tip cells will have considerably more time to accumulate synthesized calbindin than do cells in the lower part of the villus. Work carried out with non-

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laying hens and laying hens, where demand for calcium is much increased, already suggest that not all changes in calbindin levels are directly related to changes in calbindin mRNA [2, 31. It might now be interesting to re-examine these steady state models in further detail to determine how 1,25(OH),D, controls calbindin expression in response to changing physiological stimuli. Acknowledgement. Miss JCY Wu is supported by a post-graduate scholarship obtained from the British Egg Marketing Board Research and Education Trust.

References 1. Bar A, Cohen A, Edelstein S, Shemesh M, Montecuccoli G, Hurwitz S (1978) Involvement of cholecalciferol metabolism in birds in the adaptation of calcium absorption to the needs during reproduction. Comp Biochem Physiol59B :245-249 2. Bar A, Shani M, Fullmer CS, Brindak ME, Striem S (1990) Modulation of chick intestinal and renal calbinin gene expression by dietary vitamin D,, 1,25-dihydroxyvitamin D,, calcium and phosphorus. Mol Cell Endocrinol72:23-31 3. Bar A, Striem S, Mayel-Afshar S, Lawson DEM (1990) Differential regulation of calbindin-D,,, mRNA in the intestine and eggshell gland of the laying hen. J Mol Endocrinol4: 93-99 4. Clemens TL, Garrett KP, Zhou X-Y, Pike JW, Haussler MR, Dempster DW (1 988) Immunocytochemical localization of the 1,25-dihydroxyvitamin D, receptor in target cells. Endocrinology122:122&1230 5. Fullmer CS (1990) Regulation of intestinal calbindin DZsKgene expression: a solution hybridization study. Arch Biochem Biophys 283: 193-199 6. Haussler MR, Mangelsdorf DJ, Komm BS, Terpening CM, Yamaoka K, Allegretto EA, Baker AR, Shine J, McDonnell DP, Hughes M, Weigel NL, O’Malley BW, Pike JW (1989) Molecular biology of the vitamin D hormone. Recent Prog Horm Res 44:263-305 7. King IS, Paterson JYF, Peacock MA, Smith MW, Syme G (1983) Effect of diet upon enterocyte differentiation in the rat jejunum. J Physiol 344:465-481 8. Kiyama H, Wu JCY, Smith MW, Lawson EDM, Emson PC (1991) Developmental control over vitamin-D-induced calbindin gene expression during early differentiation of chicken jejunal enterocytes. Differentiation 46: 69-75 9. Lawson DEM, Wilson PW, Barker DC, Kodicek E (1969) Tsolation of chick intestinal nuclei: Effect of vitamin D3 on nuclear metabolism. Biochem J 1 15: 263-268

10. Lowe KE, Maiyar AC, Norman AW (1992) Vitamin D-niediated gene expression. Crit Rev Eukaryotic Gene Expression 2: 65109 11. Mayel-Afshar S, Lane SM. Lawson DEM (1988) Relationship between the levels of calbindin synthesis and calbindin mRNA in chick intestine. J Biol Chem 263 :4355-4361 12. Morrissey RL, Zolock DT, Bikle DD, Empson RN, Bucci TJ (1978) Intestinal response to la,25-dihydroxycholecalciferol.I. R N A polymerase, alkaline phosphatase, calcium and phosphorus uptake in vitro, and in vivo calcium transport and accumulation. Biochim Biophys Acta 538: 23-33 13. Nemere I, Norman AW (1991) Transport of calcium. In: Field M, Frizzell RA (eds) The gastrointestinal system, vol. IV. Handbook of Physiology. American Physiology Society, pp 337-360 14. Norman AW, Friedlander EJ, Henry HL (1981) Determination of the rates of synthesis and degradation of vitamin D-dependent chick intestinal and renal calcium-binding proteins. Arch Biochem Biophys 206: 305-317 15. Roche C, Bellaton C, Pansu D, Miller A, Bronner F (1986) Localization of vitamin D-dependent active Ca2 transport in rat duodenum and relation to CaBP. Am J Physiol 251 :G314(3320 16. Shinki T, Takahashi N, Kawate N, Suda T (1982) The possible role of calcium-binding protein induced by la,25-dihydroxyvitamin D 3 in the intestinal calcium transport mechanism. Endocrinology 111 : 154G1551 17. Smith MW (1991) Cell biology and molecular genetics of enterocyte differentiation. Curr Top Membranes 39: 153-179 18. Smith MW, Bruns E, Lawson EDM (1985) Identification of intestinal cells responsive to calcitriol (1,25-dihydroxycholecaIciferol). Biochem J 225 : 127-1 33 19. Smith MW, Turvey A, Freeman TC (1992) Appcarance of phloridzin-sensitive glucose transport is not controlled at mRNA level in rabbit jejunal enterocytes. Exp Physiol 77 : 525528 20. Spencer R, Charman M, Emtage JS, Lawson DEM (1976) Production and properties of vitamin D induced mRNA for calcium binding protein. Eur J Biochem 72: 399-409 21. Spielvogel AM, Farley RD, Norman AW (1972) Studies on the mechanism of action of calciferol. V. Turnover time of chick intestinal epithelial cells in relation to the intestinal action of vitamin D. Exp Cell Res 74:359-366 22. Taylor AN (1983) Intestinal vitamin D-induced calcium-binding protein : Time-course of immunocytochemical localization following 1,25-dihydroxyvitamin D,. J Histochem Cytochem 3 1:426-432 23. Theofan G, Nguyen Anh P, Norman AW (1986) Regulation of calbindin-D,,, gene expression by 1,25-dihydroxyvitamin D, is correlated to receptor occupancy. J Biol Chem 262116943-16947 +