The postnatal structural and functional changes which occur in the gut are a result of various processes, like genetic program of gut maturation, responses to dietary changes (e.g., change from milk to solid food) and recovery following injury. However, most of the studies performed so far concerned the description of the effects (e.g., mucosa histometry, DNA and protein synthesis, activity of brush-border enzymes) rather than the cellular mechanisms involved. Solving the very dynamic equilibrium between proliferation and maturation of the enterocyte as well as programmed cell death is the key to understanding what happens in the gut mucosa in macro scale. The existence of programmed cell death process in the intestine was recognized for the first time in the mid 1990 by Iwanaga (1) and Shibahara et al. (2) in adult humans and animals, while at the end of the millennium it was allocated on the entire length of the villi and in the crypts by Westcarr et al. (3). What followed was the intensive study of the extent of the apoptosis triggered by various dietary conditions or deprivation of growth factors (4-8). Interesting observations were made by Wildhaber et al. (9) who associated increased apoptosis in the enterocytes with the decrease of Bcl-2, major antiapoptotic protein, during total parenteral nutrition. Wang et al. (10) correlated apoptosis with free radicals formation during passive smoking in rats, while Xia & Talley (11) with the ongoing infection. In our previous studies "packets" of apoptotic enterocytes, i.e., neighbouring cells dying together, were observed (5, 6). This suggested the presence of auto-/paracrine factors that are involved in the promotion of cell death signal and the major role of receptor pathway of apoptosis. The TGF-ß1 was suggested as a possible cytokine responsible for the pattern, as its expression was confirmed in the mucosa of small intestine in pigs (6). Concomitantly the p53 protein, genome guard that acts via mitochondrial pathway, takes charge of enterocytes with altered DNA, especially during intensive growth and remodeling after birth and during weaning (unpublished data). The presence of programmed cell death II - autophagy, was also confirmed among enterocytes (6). On the other hand plethora of factors preventing apoptosis and/or enhancing proliferation were identified to play a role in the gut development. The most potent were colostrum and milk (12-14) together with isolated growth factors and tissue hormones (8, 15). In the meantime Blum and Baumrucker (7) reported that supplementation with the grow factors alone is not sufficient to prevent deterioration of the gut mucosa. Considering the variety of factors that induce and modulate the cell death it surprising how little is known about the mechanisms that control enterocyte cell death.

Programmed cell death

Nowadays the programmed cell death is divided into three major types. Type 1 (PCD I) consists of apoptosis and two of its derivatives: anoikis - death resulting from the detachment from lamina propria, and amorphosis - the type of cell death initiated by the distortion of the cytoskeleton. Apoptosis, the most common form of cell death, consists of two strongly intertwining pathways: mitochondrial (intrinsic) and receptor-mediated (extrinsic). In the receptor pathway death is mediated by the binding of a ligand to death receptor, that facilitates formation of death-inducing signaling complex (DISC), auto-activation of caspase 8 and by its action the activation of executor caspases (16, 17). In the mitochondrial pathway the death signal initiates hetero-oligomerization of Bax and Bid and their interaction with mitochondria membranes (18-20). This facilitates efflux of variety of factors from the intermembrane space, like cytochrome c, AIF, Smac/DIABLO, procaspases 9 and 3. Cytochrome c together with Apaf-1 forms the apoptosome where caspase 9, the regulatory caspase responsible for activation of executor caspases is auto-activated (21). The Smac/DIABLO is responsible for the inactivation of IAPs - potent inhibitors of both regulatory and executor caspases (22). The most common executor caspases for both pathways of apoptosis are caspase 3, then 6 and 7 (23), all responsible for the cleavage of the cellular structures and proteins. Caspase 8 has a potential for the activation of Bid (24) transferring signal from extrinsic to intrinsic pathway, while capase 9 directly, or by the action of caspase 3, is capable to activate caspase 8 (25). Second type of programmed cell death (PCD II) is autophagy. It was previously associated with the cell response to starvation, and is characterized by the selfdestruction of the mitochondria to minimize energy consumption. Nowadays it is believed that by entering this pathway the cell tries to avoid apoptosis by reducing the major sources of proapototic factors - mitochondria and endoplasmic reticulum (26). Only prolonged or greatly enhanced process leads to auto-destruction of the cytozol and cell death. What is interesting, that cathepsins, the major enzymes involved in autophagy are capable of activating Bax, Bid and executor caspases, which intertwines PCD II with PCD I (27-29). PCD III is the most enigmatic form of cell death where all forms of programmed cell death that does not fit to group 1 or 2 are classified.

Potential inducers of apoptosis in the gut epithelium

The genome guard, p53 protein is responsible for the detection of cells with damaged DNA. In the intestinal mucosa, especially in the early postnatal life the process od mitosis is greatly enhanced which is associated with the increasing number of DNA alterations. Upon detecting the damage, p53 blocks the cell cycle, preventing the spread of the mutation and activates DNA-repair mechanisms. If the repair is impossible p53 acts either directly, initiating mitochondrial pathway of apoptosis, or as transcription factor, via p21, enhancing the expression of proapoptotic proteins from Bcl-2 family (30-33). Both ways lead altered cell towards intrinsic pathway of apoptosis.

TGF-ß1 is a member of the transforming growth factor super-family that consists of inducers of both growth (eg. TGFalpha, TGF-ß1) (34) and death (e.g. myostatin, TGF-ß1) (35, 36). TGF-ß1 was placed deliberately in both groups because the action of this cytokine depends on the type of cell it interacts with. It is a potent inducer of cell death in mammary gland epithelium (37, 38), while it acts as a growth agonist for fibroblasts (39). It is secreted by macrophages and epithelial cells upon the uptake of apoptotic bodies (1) and its expression in the small intestine epithelium was recently confirmed (6). TGF-ß1 action is initiated by binding to the one of its receptors. Activated TGF-RIII triggers irreversible distortion in the cytoskeleton and facilitates the amorphosis in target cell. On the other hand TGF-RI and -II act via the cascade of secondary messengers, the SMAD proteins (40), which transfer the signal to the cell nucleus where they act as transfection factors, facilitating expression of various proteins, like RunX (41). RunX either directly or indirectly, via p21 (42), shifts the balance between antiapoptotic (Bcl-2) and proapototic (Bid) proteins in favor of the latter, directing the cell towards apoptosis. It is also involved in the up-regulation of Toll-like receptors in the enterocytes providing cell protection from pathogens (43). TNFalpha is a common inducer of the extrinsic pathway of apoptosis, widely expressed by leukocytes, which acts via DISC complex and caspase 8 (44).

In the present study the interactions between TGF-ß1 and TNFalpha in the molecular mechanisms of the enterocyte programmed cell death were analyzed, as well as the role of p53 and autophagy in the cell turnover in the small intestinal mucosa of newborn piglets.


MATERIAL AND METHODS
Animal preparation

All experimental procedures were approved by the Local Ethical Committee. Study was carried on 8 randomly chosen neonatal piglets aged 1 day (unsuckling neonates) and 7 days (n=4 for each age group), acquired from sows (Polish landrace x Pietrain) maintained in regular farming conditions. Sows were fed with the standard diet for pregnant (DM 87.6%, ME 11.35 MJ/kg, CP 13.1%) and lactating (DM 87.3%, ME 12.93 MJ/kg, CP 15.4%) sows. The piglets were delivered at term and healthy. Frozen (-80°C) cross-sections of middle part of jejunum (50% length) were analyzed by confocal microscopy and image analysis system.

Immunofluorescent staining

Cross section of jejunum (10 µm) mounted on silanized microscope slide (Sigma-Aldrich Chemie GmbH, Schnelldorf, Germany) were permeabilized in 70% methanol (Polskie Odczynniki Chemiczne, Gliwice, Poland), rinsed in PBS (Sigma-Aldrich Chemie GmbH, Schnelldorf, Germany), air-dried and labeled with primary antibodies (30 min in darkness). Afterwards the slides were rinsed twice in PBS and labeled with set of secondary antibodies (30 min in darkness). For triple-staining slides were afterwards stained with HOECHST 33342, incubated 10 min. After rinsing labeled slides were covered with immunofluore mounting-medium (Sigma-Aldrich Chemie GmbH, Schnelldorf, Germany) and covered with cover-glasses. Prior visualization slides were stored in +2°C in darkness. Combination of following primary anti-human antibodies was used: goat anti-TGF-RII, goat anti-TNFalpha, goat anti-Bid, goat anti-MAP I LC3, rabbit anti-TGF-ß1, mouse anti-active caspase 8, cat anti-p53 FITC-conjugated (Santa Cruz Biotechnology Inc., Santa Cruz, CA, USA) and rabbit anti-active caspase 3 (CPP32), mouse anti-Bcl-2 FITC-conjugated (DAKO, Glostrup, Denmark). The set of secondary antibodies was the folowing: Alexa Fluor 488 chicken anti-rabbit; Alexa Fluor 488 chicken anti-goat; Alexa Fluor 488 chicken anti-mouse; Alexa Fluor 546 donkey anti-goat; Alexa Fluor 568 goat anti-rabbit (Molecular Probes, Eugene, OR, USA). All primary antibodies were diluted 1:200 with PBS-1% BSA (Sigma-Aldrich Chemie GmbH, Schnelldorf, Germany), except for CPP32 and Bcl-2 which were diluted 1:100 with PBS-1% BSA. Secondary antibodies were diluted 1:500 with PBS.

Confocal colocalization studies

Cells were either double or triple stained for colocalization study on FV 500 confocal scanning microscope (Olympus Polska Sp. z o.o., Warsaw, Poland), 60x NA 1.4 oil immersion lens. For each piglet 3 cross-sections were analyzed, at least 10 images were acquired from each cross-section and representative images for each group are shown. Excitation - emission wavelengths were 488 nm vs. 505-525 nm or 543 nm vs. 610 nm. For triple staining digital, deconvolution-based "confocal" image was acquired with the use of CELL^P software on BX 40 fluorescent microscope equipped with 40x NA 0.75 air lens and motorized stage, ex. - em. filters: 405 - 480 nm, 488 - 525 nm and 543 - 560 nm (Olympus Polska Sp. z o.o., Warsaw, Poland).

Scanning cytometry

The expression of Bcl-2 and Bid in enterocytes with high and low expression of TGF-ß1 was quantitatively analyzed in 7 day old piglets with the use of SCAN^R scanning cytometer (Olympus Polska Sp. z o.o., Warsaw, Poland). Excitation - emission filters were: 405 - 480 nm, 488 - 525 nm and 543 - 610 nm.


RESULTS
TGF-ß1 in the epithelium of the small intestine of pig neonates

TGF-ß1 expression in the small intestine epithelium was already abundant at day 1 but increased even further at day 7 of life. Packets of TGF-ß1-positive cells were observed at both days (Fig. 1a - insert; b - arrows 2 and 3; Fig. 3a and b - inserts). Figure 3 shows not only the transmission of death signal via continuum in the crypt (Fig. 3b - arrow 1) but also between neighboring villi at their base (Fig. 3b - arrow 2). Expression of TGF-RII was observed on the whole circumference of the enterocyte, but it was the strongest near the basal layer (Fig. 1a - insert). Interestingly, expression of TGF-RII seamed to increase in the vicinity of cells expressing TGF-ß1 (Fig. 1a - insert and arrows). At day 1 there was no difference in the fluorescence intensity among the enterocytes expressing TGF-ß1 (Fig. 1a), but at day 7 cells varied significantly in the level of TGF-ß1 expression (Fig. 1b). The distinct pattern could be observed: cells with almost no TGF-ß1 expression, yet with high expression of TGF-RII (Fig. 1b - arrow 1), cells with intermediate, granular expression of TGF-ß1 and TGF-RII (Fig. 1b - arrow 2), finally cells with strong TGF-ß1 and TGF-RII expression located in the large packet of presumably dying enterocytes (Fig. 1b - arrow 3). Quantitative analyses showed that TGF-ß1 expression is positively associated with expression of Bid and negatively with expression of Bcl-2 (Fig. 2a). Microphotographs showed also that in TGF-ß1-positive cells the pattern of both Bcl-2 and Bid expression is granular, what is not observed in TGF-ß1-negative enterocytes, what suggests the aggregation of those proteins within the cell (Fig. 2b).

Fig. 1. Colocalization between TGF-ß1 and TGF-RII in the epithelium of middle part of jejunum in neonatal piglets. TGF-ß1 visualized by Alexa Fluor 568 (red fluorescence), TGF-RII by Alexa Fluor 488 (green fluorescence). Yellow fluorescence indicates colocalization between examined proteins. a) Day 1 of life. Insert shows expression of TGF RII on the whole circumference of the enterocytes, with the highest concentration near the basal layer. In the vicinity of cells expressing TGF-ß1 expression of TGF-RII is markedly increased (arrows, insert). b) Day 7 of life. Arrows indicate subsequent states of colocalization pattern: 1 - high expression of TGF-RII, almost no expression of TGF-ß1; 2 - high expression of TGF-RII with increased, granular expression of TGF-ß1; 3 - high expression of both TGF-RII and TGF-ß1 in the enterocytes dying together in the packet. Lens magn. 60x; insert: lens magn. 60x, 3x digital zoom.

Fig. 2. Quantitative evaluation of Bcl-2 (FITC - green fluorescence) and Bid (Alexa Fluor 488 - green fluorescence) expression in the enterocytes with high (+) and low (-) expression of TGF-ß1 (Alexa Fluor 568 - red fluorescence) in the middle part of jejunum of 7 day old piglets (a) (n=4). Panel of representative microphotographs of relocated enterocytes from each group shows a distinctive granular pattern of both Bid and Bcl-2 expression in the cells positive for TGF-ß1 (+) indicating aggregation of examined proteins on intracellular organelles occurring in the course of apoptosis (b).

Caspase 8 expression

Active caspase 8 expression is the common feature during enterocyte death. It was shown that caspase 8 colocalize with a variety of factors involved in the enterocyte programmed cell death (Fig. 3). It was strongly expressed in the packets of TGF-ß1-positive cells (Fig. 3a - inserts) and enterocytes expressing TNFalpha (Fig. 3d), but traces could be found in almost every other visible cell (Fig. 3a, c, d, f). By comparison at day 7 active caspase 8 expression was limited to those enterocytes expressing TGF-ß1 or TNFalpha (Fig. 3b and e). Colocalization of active caspase 8 with TGF-RII showed interesting feature: most of caspase 8-positive cells expressed the receptor for TGF-ß1 (as in Fig. 3c - arrow 1), there were some cells in which no expression of TGF-RII was found (Fig. 3c - arrow 2). Active caspase 8 strongly colocalized with TNFalpha at both 1 and 7 day of piglet life, but at day 7 the increase in number of enterocytes expressing TNFalpha was observed (Fig. 3 compare d and e). No clear packet pattern of cells expressing TNFalpha could be seen, but occasionally at day 7 groups of 2 or 3 of TNFalpha-positive enterocytes were present. Bid expression was abundant in enterocytes at both days and always associated with active caspase 8 (Fig. 3f).

Fig. 3. Colocalization of active caspase 8 with TGF-ß1 (a and b), TGF-RII (c), TNFalpha (d and e) and Bid (f) in the middle part of jejunum of neonatal, 1 day (a, c, d, f) and 7 day old (b and e) piglets. Active caspase 8 visualized by Alexa Fluor 488 (green fluorescence), TGF-ß1 by Alexa Fluor 568, TGF-RII, TNFa and Bid by Alexa Fluor 546 (red fluorescence). Yellow fluorescence indicates colocalization between examined proteins. Interestingly in 1 day old piglets (a, c, d, f) alongside prominent expression of active caspase 8 associated with examined proteins there is a faint expression of active caspase 8 in all enterocytes. This phenomenon disappears completely in 7 day old piglets (b and e). a and b) Clearly visible are the TGF-ß1-positive packets of dying enterocytes (inserts). Furthermore the transmission of apoptotic signal is visible in the 7 day old piglet (b) not only via continuum (arrow 1) but also via lumen between neighboring villi (arrow 2). c) Increased expression of the active caspase 8 is not always associated with high expression of TGF-RII (compare arrows 1 and 2), suggesting involvement of other cytokines as the initiators of apoptosis. d and e) TNFalpha seams to play important role in the promotion of apoptotic signal, but no clear pattern of TNFa-positive packets of enterocytes is observed. Expression of TNFalpha increases between 1 and 7 day of life. f) Colocalization between active caspase 8 and Bid may indicate transmission and amplification of apoptotic signal from the receptor to the mitochondria-mediated pathway. Lens magn. 60x; inserts: lens magn. 60x, 3x digital zoom.

Colocalization between TGF-ß1 and TNFalpha

To check whether there are interactions between TGF-ß1 and TNFalpha in the course of the programmed enterocyte death we performed the colocalization study between those cytokines expression (Fig. 4). Majority of dying enterocytes expressed both cytokines (Fig. 4 - white arrows) but there were also those expressing only one of them (Fig. 4 - red arrows for TGF-ß1, green for TNFalpha).

Fig. 4. Colocalization between TGF-ß1 (Alexa Fluor 568 - red fluorescence) and TNFalpha (Alexa Fluor 488 - green fluorescence) in the middle part of jejunum of neonatal 1 day old piglet. Cell nuclei counterstained with HOECHST 33342 (blue fluorescence). Three different patterns of cytokine expression can be observed in the enterocytes: majority expressed both cytokines (white arrows), while some only one of them (red arrows for TGF-ß1, green for TNFalpha). Image reconstructed from the stack of adjoining microphotographs by CELL^P software, lens magn. 40x.

Expression of p53 in the neonatal gut

Expression of p53 in the gut epithelium was limited strictly to the crypt region and basal part of villi (Fig. 5a - insert). Surprisingly at day 1 all of the p53-positive enterocytes coexpressed active caspase 3 - the major executor active during the late, irreversible phase of apoptosis (Fig. 5a). On day 7, by comparison, between 1/3 and half of all p53 positive cells did not show the expression of active caspase 3 (Fig. 5b). Interestingly, at day 7 in several crypts majority of enterocytes expressed p53, in contrast to day 1 when expression of p53 was evenly spread (Fig. 5 compare a and b).

Fig. 5. Colocalization between p53 (FITC - green fluorescence) and active caspase 3 (Alexa Fluor 568 - red fluorescence) in the middle part of jejunum of neonatal 1 and 7 day old piglets. Yellow fluorescence indicates colocalization between examined proteins. All p53-positive cells are localized in the crypts and on the basis of the villi (a - insert). a) In 1 day old piglet there is 100% colocalization between examined proteins which indicates that all enterocytes with DNA alterations are eliminated by apoptosis. b) In 7 day old piglet some of p53-positive enterocytes are negative for active caspase 3 suggesting repair mechanism rather than elimination (arrows). Interestingly in 7 day old piglets several crypts where all enterocytes expressed p53 were observed. Lens magn. 60x.

Autophagy in the neonatal gut

Analysis of autophagy was carried out with the use of MAP I LC3 protein, the only reliable marker of PCD II associated with formation of autophagosome membranes. At day 1 almost all enterocytes on the villi abundantly labeled with this marker (Fig. 6a - insert 2), but no such pattern was observed in the crypt area, where MAP I LC3 expression was limited to some of the cells (Fig. 6a - insert 1). Colocalization with active caspase 3 showed that while in the crypts all of MAP I LC3 cells were actually dying, on the villi the process was limited only to a few of them (Fig. 6a). Interestingly, packets of cells expressing both markers were observed in the crypts and on the villi (Fig. 6a). At day 7 the expression of MAP I LC3 colocalized almost entirely with active caspase 3 (Fig. 6b) with exception of a few enterocytes (Fig. 6b - arrow).

Fig. 6. Colocalization between MAP I LC3, considered the only reliable marker of autophagy (Alexa Fluor 488 - green fluorescence) and active caspase 3 (Alexa Fluor 568) in the middle part of jejunum of neonatal 1 and 7 day old piglets. Yellow fluorescence indicates colocalization between examined proteins. Autophagy intertwines with apoptosis, as dying cells coexpress MAP I LC3 and active caspase 3. At day 1 of life the strong pattern of MAP I LC3-related fluorescence is visible in all enterocytes on the villi (a - insert 2) while in crypts only the cells that are dying show MAP I LC3 expression (a - insert 1). The wide-ranging expression is associated with apical capillary system (ACS) involved in transport of macromolecules across the open gut barrier. It consists of vacuoles similar to those present during autophagy where MAP I LC3 is the crucial membrane element. The ACS-related expression disappeared nearly completely at day 7 (b). Thus in the neonates MAP I LC3 expression may be used as a marker of autophagy (together with active caspase 3) and a marker of gut maturation. Lens magn. 60x; inserts: lens magn. 60x, 3x digital zoom.

DISCUSSION
Analyses of molecular mechanisms of programmed cell death in the intestinal mucosa is very difficult. This complex tissue consists of a variety of different cell types, with different origin, like intestinal epithelium, connective tissue, blood vessels, Peyer patches, neurons, smooth muscles, etc. In response to single stimulus they demonstrate a plethora of, often contradictory actions. There is the need for a technique that would give the possibility to localize the observed processes within the epithelium. Confocal microscopy has been chosen because unlike other methods of analysis (i.e. western blot, real-time PCR) it gives the chance of precise allocation of the processes in the greater image of the tissue and their association with studied cell type.

The existence of packets of dying enterocytes has been reported previously (5, 6) and is presented in this publication (Fig. 1 and 3). This strongly support the idea of auto-/paracrine factors involved in the transmission of programmed cell death in the epithelium of small intestine. Our study showed that the major cytokine responsible for this pattern is TGF-ß1, which is secreted not only by the macrophages and epithelial cells upon the uptake of the apoptotic bodies (1), but at the same time it acts as a potent death inducer in the epithelial cells (37, 38). On figures 1 and 3 (a and b) the packets of enterocytes expressing significant amounts of TGF-ß1 were presented, while no such pattern was observed for TNFalpha (Fig. 3d and e). Furthermore expression of TGF-RII in the gut epithelium was abundant at early days of life (Fig. 1 and 3c), suggesting the importance of TGF-ß1-mediated pathways in the entrocyte turnover and gut maturation. Transmission of death signal was confirmed not only via mucosa continuum but also between neighboring villi (Fig. 3b), which verifies our previous observations conducted on the basis of active caspase 3 expression (6). Based on these observations we proposed the following scheme of TGF-ß1 role as the mediator of apoptotic signal between the enterocytes (Fig. 7). Upon the uptake of the remnants of dead enterocyte, the apoptotic bodies, both macrophages and neighboring cells express and secrete TGF-ß1 to their surroundings. Expression of TGF-RII on the whole circumference of the cell (Fig. 1a - insert) confirmed that signal transmission occurred not only via the basal membrane, where TGF-RII expression was strongest, but also in between the neighboring enterocytes and via the lumen. Increase in the TGF-RII expression in the vicinity of TGF-ß1-positive cells further substantiated the major role of this cytokine in the gut remodeling process (Fig. 1a - arrows). Colocalization of both TGF-ß1 and TGF-RII with active caspase 8 (Fig. 3a-c), the major regulatory caspase on the extrinsic, receptor-mediated pathway of apoptosis, suggested the grim fate of TGF-ß1-positive cells. It was surprising as there was no direct correlation between TGF-RII and DISC ever reported. We can not exclude the involvement of TGF-RIII and amorphosis (45) from playing the role in enterocyte cell death, but the abundance of TGF-RII expression in the gut epithelium strongly advocates for its supremacy. Activation of TGF-RII by its ligand lead to the cascade of secondary messengers, the SMAD proteins, which act as transcription factors facilitating expression of various proteins. Among them RunX seamed to play a key role in the enterocyte death. By its action the changes occurred in the expression of proteins from Bcl-2 family, that control apoptosis (46). It is not yet sure whether in the intestinal epithelium RunX acted alone or, as suggested by Yano et al., via p21 (42), but our study showed distinct positive correlation between expression of TGF-ß1 and Bid, while Bcl-2 was down-regulated (Fig. 2). This shifted the balance between pro- and anti-apoptotic proteins in the favor of cell death. The granular pattern of both Bid and Bcl-2 expression in TGF-ß1-positive cells (Fig. 2b) suggested aggregation of both proteins on the cell organelles, the feature common to the apoptotic process (20, 47, 48). The active caspase 8 expression however suggested that TGF-ß1 signal alone might not be strong enough to kill the cell. As there was no direct link between those two proteins there was a possibility of other cytokine interactions in the process of programmed cell death in the enterocyte. The most potent death ligands that act via DISC-related receptors were TRAIL, FasL, as well as interferons type I and type II, and the strongest of all: TNFalpha. Not surprisingly there was association between TNFa and expression of active caspase 8 (Fig. 3d-e). Strong colocalization between TGF-ß1 and TNFalpha (Fig. 4) suggested concomitance of those cytokines in the initiation phase of enterocyte apoptosis. Colocalization of active caspase 8 with Bid (Fig. 3f) may have been a result of TGF-ß1 - TNFalpha interactions, but may also imply that apoptotic signal was transmitted and amplified via the intrinsic, mitochondrial pathway of PCD I (24). Gathering together all the information we proposed that TGF-ß1, by the distortion of balance between promoters and antagonists of apoptosis from Bcl-2 family sensitized the entrocyte to the death signal mediated by other cytokines, most potent of which was the TNFalpha (Fig. 8). The marginal role of TGF-RIII-mediated amorphosis can not be abolished, but the true extent of this pathway is yet to be determined.

Fig. 7. Scheme illustrating role of TGF-ß1 as auto-/paracrine factor involved in the transmission of apoptotic signal between enterocytes. Relevant information in the text.

Fig. 8. Pathways of TGF-ß1 within the enterocyte and its concomitance with TNFalpha in the progression of apoptotic signal within the enterocyte. Further explanation in the text.

The expression of p53 was limited to the crypt region and the basis of the enterocytes, which suggested that only the dividing population of enterocytes was monitored by the genome guard (Fig. 5a - insert). Observations of Mickiewicz, Laubitz, Zabielski and Tudek (unpublished data) on the expression and localization of DNA-repair enzymes suggested that they take over the p53 functions as the major guards of DNA stability further up on the villi. Should they fail, the defective enterocyte was eliminated via apoptosis. The fate of enterocytes with DNA alterations detected by p53 seamed to be age dependant. In the 1 day old piglets, in the midst of major growth and remodeling of gut epithelium the fetal-type enterocytes were totally eliminated via apoptosis, as they all coexpressed the active form of caspase 3 (Fig. 5a). In 7 day old piglets the presence of p53-positive, caspase 3-negative enterocytes suggested that DNA repair may have occurred or simply there is a new enterocyte population lining the epithelium, since enterocyte turnover is about 3 days (Fig. 5b). Interestingly the expression of p53 was sometimes observed in all enterocytes within one crypt, some of them dying, some presumably being under repair (Fig. 5b). This strongly reminds the theory of aberrant crypts observed in the large intestine, where the carcinogenesis may start and spread from single crypt (49).

The process of autophagy is nowadays believed to be a way of cell self-defense against the apoptosis. By elimination of organelles that store large amounts of proapototic proteins, namely mitochondria and parts of endoplasmic reticulum, the cell may have prolonged its existence (29). But when overexert the process leads to full destruction of the cytosol and cell death. The only believed reliable marker of the process, the MAP I LC3 protein, proved questionable as its expression was abundant in all of the enterocytes on the villi in 1 day old piglets (Fig. 6a - compare inserts). Expression of active caspase 3 showed that only a few of those were actually dying (Fig. 6a). This uneven distribution, along with almost total disappearance from healthy enterocytes at day 7 provided the clue to understanding the phenomenon. In fetal type enterocytes at day 1 the apical capillary system (ACS) was present (4). It consisted of vacuoles, similar to those created during autophagy, with MAP I LC3 being the part of their wall. With the closure of gut barrier and disappearance of fetal-type enterocytes the expression of MAP I LC3 was once more solemnly associated with autophagy. Thus in neonatal studies there was a need to colocalize expression of MAP I LC3 with active caspase 3 to sort out the populations of dying enterocytes from those with active ACS.

Acknowledgments: Supported by grant from National Committee for Scientific Research, Poland No: PBZ-KBN-093/P06/2003 and university grant No: 504 - 02310015.


REFERENCES

1. Iwanaga T. The involvement of macrophages and lymphocytes in the apoptosis of enterocytes. Arch Histol Cytol 1995; 58: 151-159.
2. Shibahara T, Sato N, Waguri S, Iwanaga T, Nakahara A, Fukutomi H, Uchiyama Y. The fate of effete epithelial cells at the villus tips of the human small intestine. Arch Histol Cytol 1995; 58: 205-219.
3. Westcarr S, Farshori P, Wyche J, Anderson WA. Apoptosis and differentiation in the crypt-villus unit of the rat small intestine. J Submicrosc Cytol Pathol 1999; 31: 15-30.
4. Sangild PT, Fowden AL, Trahair JF. How does the foetal gastrointestinal tract develop in preparation for enteral nutrition after birth? Livestock Prod Sci 2000; 66: 141-150.
5. Zabielski R, Biernat M, Godlewski MM, Woliñski J, Motyl T. Apoptosis in the gut epithelium in neonates. Folia Univ Agric Stetin 2003; 233: 41-48.
6. Godlewski MM, S³upecka M, Woliñski J, Skrzypek T, Skrzypek H, Motyl T, Zabielski R. Into the unknown - the death pathways in the neonatal gut epithelium. J Physiol Pharmacol 2005; 56: 7-24.
7. Blum JW, Baumrucker CR. Colostral and milk insulin-like growth factors and related substances: mammary gland and neonatal (intestinal and systemic) targets. Domest Anim Endocrinol 2002; 23: 101-110.
8. Woliñski J, Biernat M, Guilloteau P, Westrom BR, Zabielski R. Exogenous leptin controls the development of the small intestine in neonatal piglets. J Endocrinol 2003; 177: 215-222.
9. Wildhaber BE, Lynn KN, Yang H, Teitelbaum DH. Total parenteral nutrition-induced apoptosis in mouse intestinal epithelium: regulation by the Bcl-2 protein family. Pediatr Surg Int 2002; 18: 570-575.
10. Wang H, Ma L, Li Y, Cho CH. Exposure to cigarette smoke increases apoptosis in the rat gastric mucosa through a reactive oxygen species-mediated and p53-independent pathway. Free Radic Biol Med 2000; 28: 1125-1131.
11. Xia HH, Talley NJ. Apoptosis in gastric epithelium induced by helicobacter pylori infection: implications in gastric carcinogenesis. Am J Gastroenterol 2001; 96: 16-26.
12. Zhang H, Malo C, Buddington RK. Suckling induces rapid intestinal growth and changes in brush border digestive functions of newborn pigs. J Nutr 1997; 127: 418-426.
13. Xu RJ, Wang F, Zhang SH. Postnatal adaptation of the gastrointestinal tract in neonatal pigs: a possible role of milk-borne growth factors. Livestock Prod Sci 2000; 66: 95-107.
14. Burrin DG, Shulman RJ, Reeds PJ, Davis TA, Gravitt KR. Porcine colostrum and milk stimulate visceral organ and skeletal muscle protein synthesis in neonatal piglets. J Nutr 1992; 122: 1205-1213.
15. Burrin DG, Wester TJ, Davis TA, Amick S, Heath J. Orally administered IGF-I increases intestinal mucosal growth in formula-fed neonatal pigs. Am J Physiol 1996; 270: 1085-1091.
16. Krammer PH. CD95's deadly mission in the immune system. Nature 2000; 407: 789-795.
17. Pajak B, Orzechowski A. FLIP - an enemy which might lose the battle against the specific inhibitors of translation. Postepy Hig Med Dosw 2005; 59: 140-149.
18. Shimizu S, Ide T, Yanagida T, Tsujimoto Y. Electrophysiological study of novel large pore formed by Bax and the Voltage-Dependent Anion Channel that is permeable to cytochrome c. J Biol Chem 2000; 275: 12321-12325.
19. Eskes R, Desagher S, Antonsson B, Martinou JC. Bid induces the oligomerization and insertion of Bax into outer mitochondrial membrane. Mol Cell Biol 2000; 20: 929-935.
20. Godlewski MM, Gajkowska B, Lamparska-Przybysz M, Motyl T. Colocalization of BAX with BID and VDAC-1 in nimesulide-induced apoptosis of human colon adenocarcinoma COLO 205 cells. Anti-Cancer Drugs 2002; 13: 1017-1029.
21. Cain K, Brown DG, Langlais C, Cohen GM. Caspase activation involves the formation of the aposome, a large (approximately 700 kDa) caspase-activating complex. J Biol Chem 1999; 274: 22686-22692.
22. Gorka M, Godlewski MM, Gajkowska B, Wojewodzka U, Motyl T. Kinetics of Smac/DIABLO release from mitochondria during apoptosis of MCF-7 breast cancer cells. Cell Biol Int 2004; 28: 741-754.
23. Nicholson DW. Caspase structure, proteolytic substrates and function during apoptotic cell death. Cell Death Differ 1999; 6: 1028-1042.
24. Kudla G, Montessuit S, Eskes R, Berrier C, Martinou JC, Ghazi A, Antonsson B. The destabilization of lipid membranes induced by the C-terminal fragment of caspase 8-cleaved Bid is inhibited by the N-terminal fragment. J Cell Biol 2000; 275: 22713-22718.
25. Slee EA, Harte MT, Kluck RM, Wolf BB, Casiano CA, Newmayer DD, Wang HG, Reed JC, Nicholson DW, Alnemri ES, Green DR, Martin SJ. Ordering of the cytochrome c-initiated caspase cascade: hierarchical activation of caspases-2, -3, -6, -7, -8 and -10 in a caspase-9-dependent manner. J Cell Biol 1999; 144: 281-292.
26. Kim J, Klionsky DJ. Autophagy, cytoplasm-to-vacuole targeting pathway, and pexophagy in yeast and mammalian cells. Ann Rev Biochem 2000; 69: 303-342.
27. Stoka V, Turk B, Schendel SL, Kim TH, Cirman T, Snipas SJ, Ellerby LM, Bredesen D, Freeze H, Abrahamson M, Bromme D, Krajewski S, Reed JC, Yin XM, Turk V, Salvesen GS. Lysosomal protease pathways to apoptosis. Cleavage of bid, not pro-caspases, is the most likely route. J Biol Chem 2001; 276: 3149-3157.
28. Bidere N, Lorenzo HK, Carmona S, Laforge M, Harper F, Dumont C, Senik A. Cathepsin D triggers Bax activation, resulting in selective apoptosis-inducing factor (AIF) relocation in T lymphocytes entering the early commitment phase to apoptosis. J Biol Chem 2003; 278: 31401-31411.
29. Yoshimori T. Autophagy: a regulated bulk degradation process inside cells. Biochem Biophys Res Comm 2004; 313: 453-458.
30. Potten CS, Merritt A, Hickman J, Hall P, Faranda A. Characterization of radiation-induced apoptosis in the small intestine and its biological implications. Int J Radiat Biol 1994; 65: 71-78.
31. Moll UM, Zaika A. Nuclear and mitochondrial apoptotic pathways of p53. FEBS Lett 2001; 493: 65-69.
32. Arima Y, Nitta M, Kunianaka S, Zhang D, Rujiwara T, Taya Y, Nakao M, Saha H. Transcriptional blockade induces p53-dependent apoptosis associated with translocation of p53 to mitochondria. J Biol Chem 2005; 280: 19166-19176.
33. Sengupta S, Harris CC. p53: Traffic cop at the crossroads of DNA repair and recombination. Nature 2005; 16: 44-55.
34. OMIM: TRANSFORMING GROWTH FACTOR, *190180
35. Motyl T, Gajkowska B, Ploszaj T, Wareski P, Skierski J, Zimowska W. Expression and subcellular redistribution of Bax during TGF-b1-induced programmed cell death of HC11 mouse mammary epithelial cells. Cell Mol Biol 2000; 46: 175-185.
36. Budasz-Swiderska M, Jank M, Motyl T. Transforming growth factor-beta1 upregulates myostatin expression in mouse C2C12 myoblasts. J Physiol Pharmacol 2005; 56: 195-214.
37. Kolek O, Gajkowska B, Godlewski MM, Motyl T. Molecular mechanism of TGF-ß1-induced apoptosis in HC11 mouse mammary epithelial cells. Cell Mol Biol 2001; 47: 197-208.
38. Kolek O, Gajkowska B, Godlewski MM, Motyl T. Antiproliferative and apoptotic effect of TGF-ß1 in bovine mammary epithelial BME UV1 cells. Comp Biochem Physiol 2003; 134: 417-430.
39. Narine K, De Wever O, Van Valckenborgh D et al. Growth factor modulation of fibroblast proliferation, differentiation, and invasion: implications for tissue valve engineering. Tissue Eng 2006; 12: 2707-2716.
40. Miyazawa K, Shinozaki M, Hara T, Furuya T, Miyazono K. Two major Smad pathways in TGF-beta superfamily signalling. Genes Cells 2002; 7: 1191-1204.
41. Chi XZ, Yang JO, Lee KY et al. RUNX3 suppresses gastric epithelial cell growth by inducing p21(WAF/Cip1) expression in cooperation with transforming growth factor b-activated SMAD. Mol Cell Biol 2005; 25: 8097-8107.
42. Yano T, Ito K, Fukamachi H et al. The RUNX3 tumor suppressor upregulates Bim in gastric epithelial cells undergoing transforming growth factor beta-induced apoptosis. Mol Cell Biol 2006; 26: 4474-4488.
43. Mikami F, Lim JH, Ishinaga H et al. The transforming growth factor-beta-Smad 3/4 signaling pathway acts as a positive regulator for TLR2 induction by bacteria via a dual mechanism involving functional cooperation with NF-kappaB and MAPK phosphatase 1-dependent negative cross-talk with p38 MAPK. J Biol Chem 2006; 281: 22397-22408.
44. Thompson CB. Apoptosis in the pathogenesis and treatment of disease. Science 1995; 267: 1456-1462.
45. Sanz-Rodriguez F, Guerrero-Esteo M, Botella LM, Banville D, Vary CPH, Bernabéu C. Endoglin regulates cytoskeletal organization through binding to ZRP-1, a member of the Lim family of proteins. J Biol Chem 2004; 279: 32858-32868.
46. Guo C, Ding J, Yao L et al. Tumor suppressor gene Runx3 sensitizes gastric cancer cells to chemotherapeutic drugs by downregulating Bcl-2, MDR-1 and MRP-1. Int J Cancer 2005; 116: 155-160.
47. Tsujimoto Y and Shimizu S. Bcl-2 family: Life-or-death switch. FEBS Letters 2000; 466: 6-10.
48. Mikhailov V, Mikhailova M, Pulkrabek DJ, Dong Z, Venkatachalam MA, Saikumar P. Bcl-2 prevents Bax oligomerization in the mitochondrial outer membrane. J Biol Chem 2001; 276: 18361-18374.
49. Luo L, Li B, Pretlow TP. DNA alterations in human aberrant crypt foci and colon cancers by random primed polymerase chain reaction. Cancer Res 2003; 63: 6166-6169.