Localization, quantification, and characterization of tuftelin in soft tissues
Article first published online: 28 MAR 2007
Copyright © 2007 Wiley-Liss, Inc.
The Anatomical Record
Volume 290, Issue 5, pages 449–454, May 2007
How to Cite
Leiser, Y., Blumenfeld, A., Haze, A., Dafni, L., Taylor, A. L., Rosenfeld, E., Fermon, E., Gruenbaum-Cohen, Y., Shay, B. and Deutsch, D. (2007), Localization, quantification, and characterization of tuftelin in soft tissues. Anat Rec, 290: 449–454. doi: 10.1002/ar.20512
- Issue published online: 13 APR 2007
- Article first published online: 28 MAR 2007
- Manuscript Accepted: 5 FEB 2007
- Manuscript Received: 10 JAN 2007
- The European Union. Grant Number: MATRIX QLK3-CT-2001-00090
- The Israel Science Foundation. Grant Number: 597/02
- The European Union. Grant Number: MATRIX QLK3-CT-2001-00090
- The Israel Science Foundation. Grant Number: 597/02
- in situ hybridization;
- soft tissues;
Tuftelin was initially found in the developing and mature extracellular enamel. Here we describe our novel discovery of tuftelin cellular distribution (protein and mRNA) in six soft tissues. The expression levels of tuftelin mRNA were significantly higher in mouse kidney and testis, in which oxygen levels are hovering closely to hypoxia under normal conditions. Anat Rec, 2007. © 2007 Wiley-Liss, Inc.
Tuftelin is an acidic protein, first discovered, mapped, and cloned from a cDNA library of ameloblasts (Deutsch et al., 1991). It is expressed by epithelial cells at very early stages of odontogenesis (Zeichner-David et al., 1997; Deutsch et al., 1998, 2002), when dentin or enamel is not yet formed. Later, when enamel formation commences, tuftelin is found in the extracellular developing enamel and concentrates at the dentino-enamel junction (DEJ), where mineralization of enamel commences (Deutsch et al., 1991, 1995). The timing of tuftelin protein expression (just before enamel mineralization commences), the location where it accumulates (at the DEJ region), and its acidic nature suggested that tuftelin might be involved in the initial stages of enamel mineralization (perhaps by nucleation) (Deutsch et al., 1991, 1998). There is also evidence that at very early stages of odontogenesis, the dental papilla mesenchyme and the preodontoblasts that eventually form the underlying dentin also express tuftelin in a transient manner (Diekwisch et al., 1997; Zeichner-David et al., 1997). Hence, it has been suggested that tuftelin might be involved in mesenchyme-ectoderm interactions during tooth development (Zeichner-David et al., 1997; Deutsch et al., 1998).
Further search of the human and mouse expressed sequence tag (EST) databases (Mao et al., 2001; Deutsch et al., 2002) and Northern blot analysis (MacDougall et al., 1998) revealed that, in addition to tuftelin expression in the developing and mineralizing enamel, partial tuftelin cDNA sequences were also detected in many normal as well as cancerous soft tissues. Initial studies indicated that tuftelin protein is also synthesized in some of these tissues, such as eye, stomach, brain, and kidney (Mao et al., 2001; Deutsch et al., 2002). However, the precise pattern of expression and localization to specific cell types within these tissues were not fully explored.
The above findings may suggest that tuftelin has a universal and/or multifunctional role and its expression is not confined to the developing and mineralizing tooth. Saarikoski et al. (2002) reported the induction of tuftelin in cancer cells during hypoxia.
In the present study, the precise cellular distribution of the tuftelin mRNA and protein was examined in six soft tissues, using immunohistochemistry coupled with confocal laser scanning microscopy, Western blot analysis, and in situ hybridization. These tissues were further examined for quantitative differences in tuftelin mRNA expression using real-time PCR. The elevated levels of tuftelin mRNA found, in the present study, in tissues that are exposed to low oxygen pressure under physiological conditions suggest a possible role of tuftelin during hypoxia.
MATERIALS AND METHODS
Animals and Tissue Preparation
Balb/c mice at ages varying from 6 weeks to 7 months were used in the present study. All experiments were approved by the Animal Care Ethical Committee, Hebrew University (Jerusalem, Israel). Immediately following dissection, tissues were fixed in PFA (paraformaldehyde (PFA) 4% for 24 hr. The tissues were then dehydrated in a series of diluted ethanol solutions, embedded in paraffin, and sectioned (5 μm). Slides were stored in the dark at −20°C to preserve antigenicity for future immunological and in situ hybridization analysis.
Slides were deparaffinized, hydrated, and rinsed, and endogenous peroxidase activity was blocked using 3% (v/v) H2O2 (diluted in methanol) for 10 min. The tissue sections were further blocked in nonimmune goat serum for 15 min at room temperature (Histostain-SP kit; Zymed, San Francisco, CA), followed by overnight incubation with the primary antibodies [polyclonal antituftelin antibodies raised against tuftelin synthetic peptides: LF74 (N-terminal region), LF75 (middle region) (Deutsch et al., 1991)], diluted in PBS, at 4°C in a humidified chamber. After rinsing, the tissue sections were treated with avidin/biotin blocking solution (Zymed, San Francisco, CA), followed by incubation with the biotinylated secondary antibody supplied by the Zymed kit. For visualization, horseradish peroxidase (HRP; Zymed) conjugate was used (brown/red precipitate). Nuclear DNA staining was performed with hematoxylin.
Confocal Laser Scanning Microscopy
The immunological procedure was similar to the immunohistochemical reaction described in the previous section. The primary antituftelin antibodies (LF-74, LF75 mix, diluted in PBS) were incubated overnight at 4°C followed by secondary antibody incubation. Two types of secondary goat-antirabbit antibodies were used: Cy5 (red) and Alexa Fluor 488 (green). Tissue sections were incubated with secondary antibodies diluted in PBS to 10 μg/ml for 2 hr at room temperature. Tissue sections were then rinsed, mounted, and examined by laser confocal microscopy (LSM 410; Zeiss, Germany). For nuclear DNA staining, propidium iodine (PI) at concentration of 1 μg/μL was used.
In Situ Hybridization
Expression of tuftelin mRNA using in situ hybridization was carried out according to manufacturer's protocol (Innogenex, San Ramon, CA). Briefly, the full mouse tuftelin sequence was RT-PCR-amplified using the following primers: DD112 (forward), 5′-CTCATACGACCTCCGCGTGAAGATG-3′, and DD108 (reverse), 5′-TCAGGTTTCCACCACCCGGATGAG-3′ (Mao et al. 2001), followed by cloning into pGEM-T Easy Vector (Promega, Madison, WI). The tuftelin sequence was amplified by PCR using SP6 and T7 primers. Antisense and sense (control) RNA probes were generated by SP6 or T7 transcription, respectively, using a DIG RNA labeling kit (SP6/T7; Roche Diagnostics, Mannheim, Germany). Probes were then treated by alkaline hydrolysis to produce short riboprobes. Hybridization of the probes was performed overnight at 40–42°C and visualized according to the standard protocol of Innogenex, using alkaline phosphatase (ALP; Innogenex) and Nitro Blue tetrazolium chloride/5-bromo-4-chloro-3′-indolyphosphate-p-toluidine salt (NBT/BCIP; Promega). Nuclear DNA staining was performed using nuclear fast red.
Real-Time Quantitative PCR
Tissues were dissected and immediately immersed in liquid nitrogen for future studies. Total RNA was extracted by homogenizing the frozen tissues in Tri-Reagent (MRC; Cincinnati, OH). RNA isolation was performed using the Tri-Reagent standard protocol. All total RNA extracts were subjected to DNase treatment (DNA-free; Ambion, Austin, TX) to eliminate any possible DNA contamination. Each sample was checked for quality on 1% agarose gel prior to reverse transcription, and the total RNA concentration was determined using the NanoDrop ND-1000 spectrophotometer (NanoDrop Technologies, Wilmington, DE). Total RNA was subjected to reverse transcription according to the manufacturer's protocol (superscript RNase H- Reverse Transcriptase kit; Invitrogene, Carlsbad, CA). Real-time quantitative PCR was performed using the LightCycler (Roche, Germany). For absolute quantification, plasmid standards were prepared. Tuftelin cDNA produced from mouse brain total RNA was cloned into a pGEM plasmid vector and subjected to DNA sequencing. The plasmid standards were diluted to yield serial dilutions of copy numbers from 1011 to 102 copies to produce a standard curve. The real-time quantitative PCR reactions were performed in a volume of 10 μl of a mixture containing the reverse-transcribed cDNA product, tuftelin-specific oligonucleotides primers (forward 5′-TGGACCCTAGCATGAGT-3′, reverse 5′-CGTTCTTGATCCGAAGC-3′, amplicon size 312 bp) and 1 μl of DNA Master SYBR green mix (Roche Molecular Biochemistry, Mannheim, Germany; including Taq DNA polymerase, reaction buffer, and dNTPs). The fluorescent signal was plotted in real-time against temperature to produce a melting curve for each sample. At the end of the run, the specificity of the reaction was determined by analysis of the PCR products on a 1–3.5% agarose gel.
RESULTS AND DISCUSSION
Expression and Localization of Tuftelin (mRNA and Protein) in Various Soft Tissues
To date, detailed distribution and localization of tuftelin protein and mRNA expression in soft tissues have not been fully explored. In the present study, we examined some of the soft tissues that were previously indicated to express partial tuftelin mRNA sequences, using blast search of EST databases (human and mouse) (Mao et al. 2001).
In the kidney, tuftelin expression was detected in the renal cortex (Fig. 1a–c) and in the outer and inner parts of the kidney medulla (Fig. 1d–e, respectively). In the cortex, the glomerulus inner part and Bowman's capsule were found to express the tuftelin protein (Fig. 1a and b) and mRNA (Fig. 1c; long arrows point to the glomerulus inner part, and short arrows to Bowman's capsule). Further evaluation of the glomerular cells expressing tuftelin, using confocal microscopy, revealed that these cells resemble morphologically either podocytes or mesangial cells (Fig. 1b, long arrow, green florescence). In the kidney medulla, the tuftelin protein was found to be expressed differently in the outer and inner renal medulla. In the outer aspect of the medulla, the expression of tuftelin was localized to the renal tubular cells (Fig. 1d). The tuftelin-positive tubular cells were of the modularly thin ascending and descending limbs of the long nefrons (red staining, arrow). A stronger and more localized reaction was detected in the endothelial vasa recta cells of the inner kidney medulla (Fig. 1e, red staining, arrow). In the retina of the mouse eye, the expression of the tuftelin protein (Fig. 1f) and mRNA (Fig. 1g) was localized to distinctive retinal layers: the ganglion cell layer (long black arrow), cells of the inner nuclear layer (short white arrow), and faintly in the rod and con layer (short black arrow). It could not be established whether tuftelin is expressed in the outer nuclear layer (asterisk). In the testis, the tuftelin protein (Fig. 1h) and mRNA (Fig. 1i) were expressed by the developing spermatocytes. Only a small amount of tuftelin exists in the spermatogonia (staining is less noticeable; short black arrow), but as spermatogenesis progresses and the spermatocytes differentiate and mature, the expression of tuftelin is increased (long black arrow). Tuftelin protein expression was also localized to Leydig cells (Fig. 1h, white arrow). In the liver, the tuftelin protein (Fig. 1j) and mRNA (Fig. 1k) were localized mainly to the liver hepatocytes cells (long arrow). Tuftelin mRNA was also detected in the endothelial layer of the blood vessels (Fig. 1k, short arrow).
Interestingly, the expression of tuftelin mRNA seems to be localized inside or in close proximity to the cell's nucleus, while the tuftelin protein seems to be localized in granular-like structures throughout the cytoplasm. In the lung, the tuftelin protein (Fig. 1l) and mRNA (Fig. 1m) were localized to the alveoli of the lung parenchyma. Tuftelin expression was localized to cells that resemble morphologically endothelial cells (black arrow). We were unable to determine whether tuftelin is also expressed by the lung pneumocytes. In the mouse brain, the tuftelin protein was localized to the brain neurons (Fig. 1n). Using confocal laser scanning microscopy (Fig. 1o), we further explored tuftelin protein expression in the mouse brain (cerebrum).
Higher magnification of the cortical/subcortical areas of the mouse brain showed that tuftelin protein expression is found almost exclusively in the neuronal cytoplasm (red staining, Fig. 1o, long arrow), but interestingly, colocalization of some tuftelin protein (red staining) and nuclear DNA (green staining, propidium iodine) was observed at few distinct and localized regions of the nucleus (yellow staining, short thick arrow), indicating the presence of tuftelin protein also in the nucleus. An indication for some tuftelin mRNA localization in the nucleus or perinuclear regions was also obtained in the mouse hepatocyte cells (Fig. 1k). Using in situ hybridization, tuftelin mRNA was localized to the pyramidal neurons of the mouse brain (Fig. 1p). Some tuftelin mRNA expression was also detected in the neuronal axon (black arrow).
Control reactions using PBS or preimmune rabbit serum replacing the first antibody were carried out for all six tissues. No staining was detected in the control immunohistochemical sections (data not shown). No staining was detected in the control in situ hybridization sections treated with the sense tuftelin probe (data not shown). Using immunohistochemistry, we were able to detect tuftelin protein expression also in the medulla of the adrenal gland, tracheal cartilage chondrocytes, mouse skin subepidermal cells, and peripheral nerves (data not shown).
Several of the tissues expressing tuftelin have a common neuronal developmental origin. These include the following. One, the developing tooth bud: teeth develop from migration of neural crest cells (Chai et al., 2000; Mitsiadis et al., 2003; Zhang et al., 2003; Hao et al., 2004; Miletich and Sharpe, 2004). Two, the retina of the eye: eyes develop from the neural plate and the optic vesicle is considered as part of the brain (Fernald, 2004; Gehring et al., 2004). Cells of the retinal ganglion cell layer, and possibly bipolar cells of the inner nuclear layer, as well as the rod and con layer, all of neuronal origin, were found to express tuftelin mRNA and protein. Three, the adrenal gland medulla, which is of neural crest origin and considered as a peripheral ganglion secreting catecholamines. Four, peripheral nerves. Five, the tuftelin protein was detected in the mouse skin (subepidermal cells; data not shown). Search of EST databases identified partial tuftelin mRNA sequences in skin melanocytes (accession no. BB756168), which also originate from neural crest. Six, tuftelin was found to be expressed in the brain, specifically in neurons. The above findings might indicate a common role for tuftelin in neuron-derived tissues.
Tuftelin expression was also detected in early stages of development. In the testis during spermatogenesis: the expression levels of tuftelin protein and mRNA increased as spermatogenesis progressed. Past and present searches of human and mouse EST databases revealed that tuftelin partial mRNA sequences were detected in as early as 4 cell stage of embryogenesis in embryonic stem cells, and in embryonic stem cells differentiated into an early endodermal cell type (accession no. CN720080, AU045470, CN428852, DR158860, and CX164695, respectively). Tuftelin was previously found to be expressed during early stages (as early as E13) of mouse embryonic tooth bud formation (Zeichner-David et al., 1997). The expression of tuftelin during the development and maturation processes of several of the above tissues implies that tuftelin might have a possible role in the developmental processes activated during organogenesis.
Western blot analysis of tuftelin protein expression in the six tissues examined in the present study (eye, brain, lung, liver, kidney, and testis; Fig. 2) revealed that the tuftelin protein is expressed in all these tissues; however, the size distribution of tuftelin polypeptides varied. All tissues expressed the 64 KDa tuftelin protein, but at various degrees. This polypeptide was the major peptide detected in the lung, liver, kidney, and testis (Fig. 2, lanes 3, 4, 5, 6, respectively), but some of the tissues also expressed tuftelin proteins of other molecular weights. In the eye (Fig. 2, lane 1), the strongest tuftelin bands were detected as 32, 30, and about 20 KDa protein bands, and additional but fainter tuftelin bands were detected in the range of 64 down to 32 KDa. In the brain (Fig. 2, lane 2), the strongest tuftelin bands were detected at about 55 to 45 KDa protein bands, but fainter higher and lower tuftelin bands were also detected. The differences in molecular weights of the tuftelin polypeptides might represent degradation products, but some may also represent tissue-specific alternative splices, which might have different functions in the different tissues examined. Tuftelin mRNA expression in the six mouse soft tissues at normal physiological conditions was quantified using the real-time quantitative PCR (Fig. 3). Results are presented as mRNA copy number per 1 microgram of total high-quality RNA (RNA quality was measured after DNase treatment, by 260/280 nm ratio using the NanoDrop spectrophotometer). Bustin (2000) and later Tricarico et al. (2002) did not find an appropriate reference gene to be used when several tissue biopsies are compared and advocated normalization to total RNA as the appropriate method of comparing steady-state mRNA levels between different in vivo tissue biopsies. This approach avoids the controversies and validation of expression of housekeeping reference genes between the tissues tested. The disadvantage of this approach is the relatively large amounts of high-quality RNA required from each tissue tested (Dheda et al., 2004).
At first, any statistically significant difference between all six groups was determined using the Kruskal-Wallis test (nonparametric ANOVA). The result was significant (P = 0.001). In order to determine significant differences in expression, we divided the results into two groups: the higher tuftelin expressing group, which included the eye, kidney, and testis, and the lower tuftelin expressing group, which included the liver, lung, and brain. Using the Mann-Whitney test and the Bonferroni correction for multiple comparisons, we compared all tissues to the liver, the highest expressing tissue from the lower expressing group. The expression of tuftelin was found to be significantly higher in the testis and kidney (P < 0.016 and < 0.008, respectively), while tuftelin expression in the eye was not significantly different. Tuftelin expression in the brain was significantly lower (P < 0.008), while the expression of tuftelin in the lung was not significantly different. The different levels of tuftelin mRNA expression between tissues could partially be explained from results published by Saarikoski et al. (2002). He showed, using representational difference analysis, three novel hypoxia-inducible genes: MIG-6, adipophilin, and tuftelin. During normal physiological conditions, both testis and kidney tissues are exposed to low oxygen pressures hovering closely to hypoxia (Aukland and Krog, 1960; Baumgartl et al., 1972; Epstein et al., 1987; Silva, 1987; Max, 1992; Brezis et al., 1994; Wenger and Gassmann, 1997; Depping et al., 2004; Neuhofer and Beck, 2005). Evidence of low oxygen pressures has also been reported during formation and mineralization of hard tissues such as bone and cartilage (Lennon et al., 2001; Schipani et al., 2001). Further studies need to be conducted to establish whether tuftelin is involved in the compensatory cycles that are triggered during hypoxia, which help different cells to adapt and survive changes in environmental conditions such as hypoxia.
Future characterization of the tuftelin precise function(s) may contribute to the understanding of tuftelin biological role(s) during hypoxia at physiological conditions, at induced hypoxia, during cancer progression, and also during biomineralization of hard tissues.
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