In Vivo Expression of Toll-Like Receptor 2 and 4 by Renal Epithelial Cells: IFN-γ and TNF-α Mediated Up-Regulation During Inflammation¹

At least two members of the Toll-like receptor (TLR)³ family are involved in the innate defense mechanism against bacterial infections (1, 2). TLR4 recognizes Gram-negative bacteria via the LPS moiety that is present in the surface of these microorganisms (3, 4, 5). Another member of this receptor family, TLR2, induces responsiveness to bacterial lipoproteins and components of Gram-positive bacteria such as peptidoglycan, indicating the involvement of TLR2 in the resolution of infections (6, 7, 8, 9). After stimulation by bacterial products, both receptors trigger the cell to produce inflammatory mediators. This process is mediated via a MyD88-dependent intracellular signaling pathway that causes translocation of NF-κB. The latter induces the transcription of the genes encoding for cytokines, chemokines, and adhesion molecules crucial to the inflammatory process aimed at clearance of invading bacteria (1, 10).

Both TLR2 and TLR4 are predominantly expressed by monocytes/macrophages and neutrophils (11). Lower expression of TLR is observed in vitro by several other leukocytes, endothelial cells, epithelial cells, and fibroblasts (12, 13, 14, 15, 16). In vitro experiments indicate that TLR expression is modulated by bacterial products and cytokines (11). Accordingly, an IFN-γ-responsive element is found in the promoter region of the gene encoding for TLR4 (17). Although the above indicates putative pathways that regulate TLR expression, the factors that mediate TLR expression in vivo remain to be elucidated.

Functional TLR4 expression is required for the resistance to experimental pyelonephritis induced by a bladder inoculum with Gram-negative bacteria. This is evidenced by the persistence of Gram-negative bacteria in the kidney after pyelonephritis induction in C3H/HeJ mice (18) that carry a malfunctioning TLR4 (3), while mice bearing intact TLR4 rapidly clear bacteria from the urinary tract. The cellular origin of TLR expression in the kidney responsible for resistance to bacteria is unknown. Considering that the kidney is an important port of entry for bacteria, we set out to localize TLR expression in the kidney and to study the role of cytokines in the regulation of TLR expression in vivo. We used a renal ischemia/reperfusion (I/R) model that leads to transient tissue damage that is associated with an inflammatory process that develops rapidly during reperfusion (19). The enhanced expression of macrophage-inflammatory protein-2, KC, TNF-α, IFN-γ, and MHC class I and II molecules in this renal sterile inflammation model (20, 21, 22) resembles the pattern of production of immunological mediators after bacterial infections to a large extent. This model enabled us to study the modulation of the expression of TLR2 and TLR4 mRNA by cytokines in vivo during inflammation.

The following Abs were used: anti-murine IFN-γ mAb F3 and anti-murine TNF-α mAb TN3 were kindly provided by HBT (Uden, The Netherlands) and Celltech (Slough, U.K.) respectively; polyclonal rabbit anti-murine TLR4 serum was kindly provided by B. Beutler (The Scripps Research Institute, La Jolla, CA); peroxidase-conjugated goat anti-rat IgG and peroxidase-conjugated goat anti-rabbit IgG were from Jackson ImmunoResearch Laboratories (West Grove, PA); anti-Tamm Horsfall protein was from Cappel (Durham, NC); sheep anti-digoxigenin/alkaline phosphatase was from Roche (Basel, Switzerland); and biotinylated rabbit anti-goat IgG was from DAKO (Glostrup, Denmark). Other reagents were proteinase K (Life Technologies, Paisley, U.K.); dextran sulfate (Pharmacia, Uppsala, Sweden); streptABComplex/AP (DAKO); polyvinylalcohol, formamide, and 2-ME (Merck, Darmstadt, Germany); DTT and salmon sperm DNA (Sigma-Aldrich, St. Louis, MO); and T7 and SP6 RNA polymerase, tRNA, nitroblue tetrazolium, and bicholylindolyl phosphate (Boehringer Mannheim, Mannheim, Germany).

All experiments were approved by the Institutional Animal Care Committee of the University of Maastricht (Maastricht, The Netherlands). Renal ischemia was induced as described (20, 21). In brief, male Swiss mice (Charles River Breeding Laboratories, Heidelberg, Germany) were anesthetized and body temperature was maintained at 39°C. After laparotomy, ischemia was induced by clamping the left renal pedicle for 45 min, during which the wound was covered. Subsequent to removal of the clamp, the contralateral kidney was removed and stored. After closing the abdomen, mice were supplemented with prewarmed PBS to maintain fluid balance. The animals were sacrificed at 1 and 6 h as well as 1, 3, and 5 days after ischemia. The experimental left kidney was harvested immediately and divided into specimens for assays described below. Mice subjected to ischemia were treated 10 min before reperfusion with 0.5 ml of PBS i.p. containing 300 μg of anti-IFN-γ mAb F3, 1 mg of anti-TNF-α mAb TN3, or PBS only. Previously, we have shown that mice receiving an isotype-matched control Ab did not differ in inflammatory parameters or renal injury when compared with the ischemia/PBS control group (20, 21), indicating the specific effects of the respective Abs.

Oligonucleotide primers were designed for the specific PCR amplification of a fragment of murine TLR2 and TLR4. The TLR2 primers were 5′-TCT GGG CAG TCT TGA ACA TTT-3′ (sense primer) and 5′-AGA GTC AGG TGA TGG ATG TCG-3′ (antisense primer), yielding a 321-bp fragment. The TLR4 primers were 5′-GCA ATG TCT CTG GCA GGT GTA-3′ (sense primer) and 5′-CAA GGG ATA AGA ACG CTG AGA-3′ (antisense primer), yielding a product of 406 bp (all primers were synthesized by Eurogentec, Seraing, Belgium).

TLR2 and TLR4 cDNA fragments were amplified from murine kidney cDNA prepared from total RNA by reverse transcription, as described below. The fragments were both TA cloned into pGEM-Teasy (Promega, Madison, WI). All clones obtained contained the correct TLR sequences that were evaluated using the Big Dye termination cycle sequencing kit (PerkinElmer/Cetus, Emeryville, CA), according to the manufacturer’s instructions. DH5-α-competent cells were transformed by heat-shock procedures. Subsequently, clones containing plasmids with the TLR insert were selected, isolated, and purified using the Qiafilter Plasmid midi kit (Qiagen, Hilden, Germany). Sense probes were prepared after linearization of the plasmid with NcoI and transcription with SP6 polymerase; antisense probes were prepared from the plasmid after linearization with SpeI by transcription with T7 polymerase. cRNA probes were labeled by incorporation of digoxigenin-labeled UTP following the manufacturer’s protocol (Boehringer Mannheim). In situ hybridization was performed as described by de Boer et al. (23) using 3-μm paraffin sections placed on coated slides (SuperFrost Plus; Menzel-Gläser, Braunschweig, Germany). Briefly, after prewarming the sections and subsequent rehydration with a decreasing xylene and ethanol gradient, the sections were hybridized with 30 ng probe in 300 μl for 16 h. The hybridization was performed at 50°C for TLR2 and at 55°C for TLR4 using a solution containing 50% formamide, 1 mg/ml tRNA, 10% dextran sulfate, 10 mM DTT, 0.25 mg/ml salmon sperm DNA, and 4× SSC. Subsequently, sections were first washed in 2× SSC with 50% formamide at 37°C, second in 0.1× SSC with 20 mM 2-ME at 42°C, and, finally, sections were treated with 100 U/ml RNase T1 in 2× SSC with 1 mM EDTA at 37°C. Digoxigenin-labeled hybrids were detected with alkaline phosphatase-conjugated sheep anti-digoxigenin, using nitroblue tetrazolium as chromogen and bicholylindolyl phosphate as coupling agent. Polyvinylalcohol was used to enhance the signal.

For RT-PCR, total RNA was extracted from kidneys using the SV Total RNA isolation system (Promega) and treated with RQ1 RNase-Free DNase (Promega). Total RNA was reverse transcribed using oligo(dT) primer and Moloney murine leukemia virus reverse transcriptase (Life Technologies) according to the supplier’s recommendations. cDNA samples were standardized based on the content of β-actin cDNA as housekeeping gene. β-actin cDNA was evaluated by performance of a β-actin PCR on multiple dilutions of each cDNA sample. The amount of amplified product was estimated by densitometry of ethidium bromide-stained 1.2% agarose gels using a CCD camera and Imagemaster VDS software (Pharmacia). The TLR primers used for the generation of the cRNA probes were also used for PCR amplification. Primers for murine β-actin were 5′-TAA AAC GCA GCT CAG TAA CAG TCG G-3′ (sense primer) and 5′-TGC AAT CCT GTG GCA TCC ATG AAA C-3′ (antisense primer); primers used for the amplification of murine TNF-α mRNA, 5′-GGC AGG TCT ACT TTG GAG TCA TTG C-3′ and 5′-ACA TTC GAG GCT CCA GTG AAT TCG G-3′ (antisense primer); primers designed for amplification of murine IFN-γ mRNA, 5′-AGC GGC TGA CTG AAC TCA GAT TGT AG-3′ (sense primer) and 5′-GTC ACA GTT TTC AGC TGT ATA GGG-3′ (antisense primer). All primers were used for PCR amplification of murine cDNA kidney samples from mice exposed to the interventions. PCR with TLR2-, TLR4-, β-actin-, IFN-γ-, or TNF-α-specific primers were performed using appropriate dilutions of the cDNA. PCR were performed in a total volume of 25 μl in PCR buffer (PerkinElmer/Cetus) in the presence of 0.2 mM dNTP (Pharmacia), 1 μM of each primer, 0.3 mM MgCl₂, and 0.5 U of Taq polymerase (PerkinElmer/Cetus). PCR conditions for each primer couple were as follows: β-actin, 95°C for 30 s, 60°C for 45 s, and 72°C for 30 s during 21 cycles; TLR2, 95°C for 30 s, 57°C for 30 s, and 72°C for 45 s during 33 cycles; TLR4, 95°C for 45 s, 61°C for 45 s, and 72°C for 45 s during 36 cycles; IFN-γ, 95°C for 30 s, 63°C for 30 s, and 72°C for 30 s during 40 cycles; TNF-α, 95°C for 30 s, 63°C for 45 s, and 72°C for 30 s during 38 cycles. Levels of TLR2, TLR4, TNF-α, and IFN-γ RNA expression were evaluated by densitometric image analysis, as described above. Relative TLR mRNA levels were calculated by comparison of band intensities of the TLR RT-PCR products with standard curves prepared by PCR amplifications on dilution series of a highly concentrated murine kidney cDNA.

Staining with the anti-Tamm Horsfall protein on paraffin sections was performed to discriminate between distal tubules, proximal tubular epithelium, the loop of Henle, and collecting ducts. Immunohistochemistry was conducted on sections adjacent to those used for in situ hybridization. Paraffin sections treated as described above were incubated with appropriate dilutions of primary Ab, washed, developed using biotinylated rabbit anti-goat IgG and alkaline phosphatase-labeled strepABComplex, and visualized, as described above.

Renal tissue samples for evaluation of TLR4 protein expression were obtained from mice of which the renal blood vessels were flushed, immediately after the mice were sacrificed, with ice-cold PBS containing nitroprusside and heparin by canulation of the left ventricle and opening of the vena cava. This was done to remove TLR4-expressing blood leukocytes. Western blotting on renal tissue was performed using 8% polyacrylamide SDS gels. Aliquots (50 μl) of kidney homogenates (50 mg/ml) in 2% SDS sample buffer were subjected to SDS-PAGE and transferred to polyvinylidene fluoride membranes (Immobilon P; Millipore, Bedford, MA). After transfer of the proteins, membranes were blocked with 5% nonfat dry milk in 50 mM Tris, 150 mM NaCl, 0.1% Tween 20, pH 7.4 (TBST). Membranes were then incubated with polyclonal rabbit anti-murine TLR4 serum (24), kindly provided by B. Beutler (The Scripps Research Institute), at a 1/2500 dilution in TBST with 0.5% nonfat dry milk. Positive bands were visualized by chemiluminescence technology (Supersignal; Pierce, Rockford, IL) using peroxidase-conjugated goat anti-rabbit IgG at a 1/5000 dilution.

To gain insight into renal expression of TLR, we evaluated mRNA expression of both TLR2 and TLR4 by RT-PCR in cDNA samples of healthy kidneys. Constitutive expression of both renal TLR2 and TLR4 mRNA was observed (Fig. 1). To determine the cellular origin of this constitutive expression, TLR mRNA was localized by in situ hybridization. As depicted in Figs. 2, A and B, and 3, A and B, basal expression of TLR2 and TLR4 mRNA was observed predominantly in the epithelial cells of the distal and proximal tubules and in Bowman’s capsular epithelium. Expression in glomeruli and endothelial cells was minor. While macrophages are reported to express relatively abundant levels of TLR2 and TLR4 mRNA, it appeared that epithelial cells express the majority of the TLR mRNA in the kidney. Only few resident macrophage-like cell types were observed in the renal tissue, and these cells stained to a similar extent as tubular epithelium. Proximal epithelial cells in these healthy kidneys stained slightly more intensely for TLR4 mRNA compared with the distal epithelium. Control incubations using sense riboprobes were negative for healthy and experimental kidneys (Fig. 3 G).

Mice were exposed to unilateral renal I/R, a process that leads to tissue damage and a subsequent sterile inflammatory reaction. This inflammatory process is characterized by initial tubular apoptosis, followed by KC, macrophage-inflammatory protein-2, and TNF-α expression and subsequent neutrophil influx with apoptosis, necrosis, and organ dysfunction peaking at day 1 (19, 20, 22). Thereafter, a strongly enhanced IFN-γ-mediated MHC-I and II expression occurs from day 3 onward (21, 25). The effect of this sterile inflammatory process on TLR2 and TLR4 mRNA levels was assessed by RT-PCR analysis. Levels of renal TLR2 mRNA decreased to undetectable levels in animals sacrificed at 1 and 6 h after ischemia. TLR2 mRNA returned to basal level after 1 day (Fig. 1). A similar mRNA expression pattern was observed for TLR4 (Fig. 1): TLR4 mRNA expression was undetectable at 1 h; from 1 to 6 h, the TLR4 synthesis returned to detectable levels. After 24 h of reperfusion, TLR4 expression was enhanced when compared with controls. TLR2 and TLR4 mRNA expression were similar in healthy and sham-operated controls, indicating that the surgical procedure did not influence TLR expression (data not shown). The results of densitometric quantification of TLR2 and TLR4 mRNA levels at 1, 6, and 24 h after ischemia are depicted in Fig. 4, A and B. At days 3 and 5 after renal ischemia, a significant increase of both TLR2 and TLR4 mRNA was detected in the kidney (Fig. 1). A 4- to 5-fold increase of TLR2 mRNA was observed when compared with healthy controls (Fig. 5,A). Similarly, a 4- and 5.5-fold enhancement of TLR4 mRNA was measured at 3 and 5 days postischemia, respectively (Fig. 5 B).

To elucidate the cellular origin of the described enhanced TLR2 and TLR4 expression in this renal inflammation model, TLR mRNA was localized by in situ hybridization. Histology showed that renal injury was clearly present 1 day after ischemia, which is associated with the influx of neutrophils (20, 25). In this inflamed tissue, reduced TLR2 mRNA expression was observed in the cortex compared with healthy controls (Fig. 2,D). In the medulla, TLR2 transcripts appeared up-regulated in the thin limb of Henle’s loop (Fig. 2,C). Moreover, TLR2 transcription was detected in macrophage-like cells in the interstitial tissue (Fig. 2, C and D). One day after ischemia, TLR4 expression was predominantly observed in the epithelial cells of the distal tubules, the thin limb of Henle’s loop, and collecting ducts, while low expression was observed in the proximal tubules, glomeruli, and capsular epithelium. A small number of macrophage-like cells were present and stained positive. In concordance to the enhanced TLR mRNA expression as measured by RT-PCR at 5 days after ischemia, intense staining for both TLR2 and TLR4 mRNA was detected by in situ hybridization at the same time point. The intense staining was predominantly localized in the collecting ducts, the thin limb of Henle’s loop, and distal tubules (Figs. 2, E and F, and 3, E and F). Identification of distal tubules in these experiments was performed by Tamm Horsfall protein staining (Fig. 3,H). At 5 days postischemia, a small number of macrophage-like cells were present that stained positive for TLR2 and TLR4. However, compared with the staining in collecting ducts, the loop of Henle, and distal tubules, this TLR expression by macrophage-like cells constituted only a minority of the total expression (Figs. 2, E and F, and 3, E and F). At day 5, only moderate TLR expression was detected in the glomeruli and proximal tubules (Figs. 2, E and F, and 3, E and F). Taken together, renal I/R leads to marked enhancement of both TLR2 and TLR4 mRNA levels, which appeared to be predominantly present in epithelial cells of the distal tubules, the collecting ducts, and Henle’s thin loop.

Renal I/R results in TNF-α-modulated renal inflammation that is followed by IFN-γ-dependent up-regulation of MHC class I and II molecules, indicating an active cytokine-mediated immune response in the organ (20, 21, 25). The role of these cytokines in the observed up-regulation of TLR mRNA expression in our I/R-induced renal inflammation model was evaluated by blocking TNF-α and IFN-γ with inhibitory Abs. The observed enhancement of renal TLR2 mRNA expression 5 days after ischemia was completely prevented in animals treated with anti-IFN-γ or anti-TNF-α Abs (Fig. 6, A and B). Moreover, blocking of either IFN-γ or TNF-α reduced the TLR2 mRNA expression at day 5 to below the constitutive level (Fig. 6,B). In contrast to this major inhibitory effect of anti-TNF-α on TLR2 mRNA synthesis in our renal inflammation model, blocking of TNF-α resulted only in a partial reduction of TLR4 mRNA expression (Fig. 6, A and C). Similar to the effect of anti-IFN-γ on TLR2 mRNA expression, blocking of IFN-γ completely prevented the enhancement of TLR4 mRNA synthesis at 5 days after ischemia, resulting in TLR4 mRNA expression below the constitutive level. Our findings indicate that TLR2 and TLR4 expression are regulated differentially by TNF-α and IFN-γ. Because both IFN-γ and TNF-α are involved in the enhanced TLR mRNA expression, we determined the effect of cytokine inhibition on cytokine mRNA expression. Anti-IFN-γ treatment decreased IFN-γ mRNA translation in the kidneys 5 days after the ischemic insult, whereas TNF-α mRNA was not decreased by inhibition of IFN-γ (Fig. 7,A). Anti-TNF-α Ab administration did not decrease the renal TNF-α and renal IFN-γ mRNA levels when compared with PBS-treated animals (Fig. 7 B). These findings show that IFN-γ does not enhance TLR mRNA expression in our model by induction of TNF-α and that TNF-α does not enhance TLR mRNA expression by induction of IFN-γ.

Renal TLR4 expression was evaluated by Western blotting using specific anti-murine TLR4 antiserum. Whole renal tissue samples, and liver as positive control, were homogenized in SDS, and proteins were separated by SDS-PAGE. Subsequent immunoblotting with the anti-murine TLR4 antiserum revealed positive bands with an apparent molecular mass of ∼95 and ∼120 kDa in normal liver and in kidney tissue 5 days after renal I/R (Fig. 8). The calculated molecular mass of mature TLR4 based on its amino acid sequence is 93.5 kDa and is consistent with the anti-TLR4-positive bands at ∼95 kDa in the Western blot. The band at a molecular mass of ∼120 kDa in renal tissue after I/R and in normal liver probably represents a glycosylated form of the TLR4 protein. The observed molecular mass form of 120 kDa of murine TLR4 is consistent with the reported apparent molecular mass of human rTLR4 (26). Basal TLR4 protein expression was not detectable with the used Western blotting technique in healthy murine renal tissue (Fig. 8). These observations show that the up-regulation of TLR4 mRNA expression during renal inflammation induced by I/R is accompanied by a significant increased TLR4 protein expression in the kidney.

In this work, we describe the mRNA localization for TLR2, the apparent receptor for products of Gram-positive bacteria (27), and for TLR4, which recognizes LPS present on Gram-negative bacteria (3), in renal tubular epithelial cells. Our description of TLR2 and TLR4 mRNA localization in the kidney is the first to show constitutive TLR2 and TLR4 in vivo expression in the epithelial cells of the proximal and distal tubules. Moreover, we observed that a nonmicrobial inflammation as induced by I/R causes remarkably enhanced TLR2 and TLR4 mRNA expression by epithelial cells of the distal tubules, the thin limb of the loops of Henle, and collecting ducts. The enhanced TLR mRNA expression during renal inflammation was found to be mediated by both IFN-γ and TNF-α and is associated with a major increase of renal TLR4 protein expression. While in vitro TLR4 mRNA expression and responsiveness had been reported for several cell types such as leukocytes, endothelial cells, epithelial cells, and fibroblasts, but most abundantly for macrophages (11, 12, 13, 14, 15), the differential expression of TLR4 in vivo remained to be elucidated. Our study indicates that murine renal epithelial cells express TLR2 and TLR4 in vivo. The observation that TLR2 and TLR4 mRNA expression in the kidney is predominantly located in renal tubular epithelium appears to have some implications for the understanding of the innate immune defense mechanism in the kidney against pyelonephritis as a result of ascending urinary tract infections (UTI). Gut-derived Gram-negative and Gram-positive bacteria are the most prevalent microbial pathogens responsible for UTI, with Escherichia coli being the most common (28). Our data suggest that tubular epithelial cells can monitor the presence of both types of bacteria. In this context, Hagberg et al. (18) showed already in 1984 that C3H/HeJ mice, who lack functional TLR4 (3), are highly susceptible to persistent Gram-negative pyelonephritis. TLR-dependent cellular activation leads to the translocation of NF-κB to the nucleus, which leads to transcription of genes encoding for cytokines, chemokines, adhesion molecules, and antimicrobial peptides (1, 3, 10, 29). Consistently, TLR4 signaling, induction of CXC chemokine expression and CXCR signaling (18, 30), and subsequent neutrophil recruitment are crucial for the clearance of Gram-negative bacteria from the kidney (31, 32). Our results suggest that the functional role of TLR4 in prevention of pyelonephritis as described (18) could be the result of functional TLR4 expression by the renal tubular epithelial cells. To monitor bacteria in the lumen of tubules and collecting ducts, the TLRs should be expressed on the apical membrane of the renal epithelial cells. It should be noted that we located renal TLR expression at the mRNA level, and obviously these results need to be confirmed by studies at the protein level by immunohistochemistry and with functional studies. In regard of the functionality of TLR4 in the kidney, it is noteworthy that the mRNA for MD-2, the essential cofactor for TLR4, is abundantly present in the murine kidney (3). We have confirmed the abundant renal MD-2 mRNA expression by RT-PCR analysis, and also observed elevation of MD-2 mRNA after I/R (data not shown).

We applied a renal I/R model to elucidate the regulation of TLR expression during inflammation. This model allowed us to evaluate cytokine-mediated TLR mRNA expression in an inflammatory process that is not influenced by bacterial products. Most interestingly, a shift occurs from the observed diffuse proximal and distal staining for TLR2 and TLR4 mRNA in healthy kidneys to a predominant and enhanced expression 5 days after ischemia in the distal epithelial tubular cells, the thin limb of the loops of Henle, and collecting ducts. Following renal I/R injury, monocyte and T cell influx has been reported from day 3 onward (33). We observed a small number of macrophage-like cells that stained all positive for both TLR2 and TLR4; however, compared with the enhanced expression in collecting ducts, the thin limb of the loops of Henle, and distal tubules, this constituted only a minority of the total TLR expression. The increased TLR2 and TLR4 mRNA expression in our renal inflammation model by epithelial cells in the distal part of the nephron and the medulla suggests the mobilization of TLR-dependent antimicrobial potential during inflammation to the site in which ascending bacteria may enter the kidney.

The I/R-induced TLR mRNA expression in the later phase of reperfusion (day 3–5) was dependent on the action of IFN-γ and TNF-α. Previously, we demonstrated that anti-TNF-α reduces kidney neutrophil influx and deterioration of renal function in our murine model of renal I/R (20), whereas anti-IFN-γ did not affect these parameters (21). The inhibition of TLR up-regulation by anti-TNF-α is not caused by a major effect on the IFN-γ mRNA synthesis (Fig. 7). This observation is supported by data showing that anti-TNF-α does not block the up-regulation of MHC molecules in this model, while this process is largely IFN-γ dependent (20, 25). Because blocking of either TNF-α or IFN-γ results in inhibition of the majority of TLR2 expression, it appears that the observed enhancement of TLR2 expression is elicited by a synergistic action of these cytokines. TLR4 synthesis is only partially blocked by anti-TNF-α and almost completely by anti-IFN-γ; therefore, it appears that the up-regulation of TLR4 in our inflammation model is primarily mediated by IFN-γ. Thus, the regulation of TLR4 mRNA expression discerns with TLR2 in our in vivo model in respect to the dependence on TNF-α. Consistently, also in vitro studies indicate that TLR2 and TLR4 expression are regulated via distinct pathways (11). Our in vivo observation of IFN-γ-mediated TLR4 expression is in line with the presence of a functional IFN response factor motif in the promoter region of the human and mouse TLR4 gene (17). It should be mentioned that we cannot exclude the possibility that TNF-α or IFN-γ acts indirectly by stimulation of a putative modulator of TLR expression.

In general, IFN-γ is involved in bacterial clearance during the late stages of infection with virulent Gram-negative or Gram-positive bacteria (34, 35, 36). Furthermore, IFN-γ deficiency is found to be associated with infection by poorly pathogenic Mycobacterium strains and Salmonella (37). Our observations suggest that IFN-γ augments enhanced reactivity to bacteria in vivo by up-regulation of TLR2 and TLR4 mRNA. Indeed, increased expression of TLR4 by IFN-γ may explain the enhanced LPS-induced lethality of IFN-γ-treated rabbits and the enhanced LPS sensitivity of monocytes/macrophages induced by IFN-γ (38, 39, 40). It appears that increased TLR2 and TLR4 expression may play a role in the mechanisms involved in IFN-γ-mediated resistance to virulent bacteria.

Although an obvious role for IFN-γ in UTI in humans is not reported, it must be mentioned that the epithelial cells of the kidney stain strongly positive with anti-IFN-γ receptor Abs in immunohistochemistry (41), indicating the responsiveness of these cells to IFN-γ. IFN-γ deficiency was observed to increase the susceptibility of mice to experimental UTI with uropathogenic E. coli (42). The latter is in line with the IFN-γ-mediated TLR4 expression (this study) and increased susceptibility of TLR4-deficient mice for Gram-negative UTI (18). Next to behavior and anatomical impairments of the urinary tract, genetic factors also seem to be involved in human ascending UTI (27). However, the genetic influence in human UTI needs further investigation (see Ref. 30).

In conclusion, TLR2 and TLR4 are constitutively expressed in both proximal and distal tubular renal epithelial cells in vivo. During renal inflammation, TLR2 and TLR4 mRNA synthesis is enhanced by the action of TNF-α and IFN-γ, and this increased expression is mainly localized in distal tubules, the thin limb of the loops of Henle, and collecting ducts. The epithelial localization of TLR mRNA expression suggests a role for epithelial-derived TLR signaling in the inflammatory response observed during ascending UTI.

We thank Dr. B. Beutler (The Scripps Research Institute) for kindly providing the anti-murine TLR4 Ab. We thank Dr. D. Rupa from the Department of Pathology, University Hospital Maastricht, and J. J. Baelde from the Department of Pathology, Leiden University Medical Center, for expert technical assistance.