Abstract We previously reported a substantial correlation between serum parathyroid hormone (PTH) levels and the myeloma response to proteasome inhibition that suggests a crucial role for the PTH receptor 1 system in the control of myeloma tumor growth. While investigating the role of PTH in the antimyeloma effect, we observed the recovery of serum PTH levels after thyroparathyroidectomy (TPTX). Although the presence of thymus-derived PTH has been reported previously, the existence or role of thymic PTH in the serum remains controversial. Here, TPTX was performed in 8- to 12-week-old C57BL/KaLwRij mice to delineate the potential source(s) for the recovery of serum PTH. Immediately after TPTX, the expected loss of measurable serum PTH was observed. Serum PTH levels recovered 3 to 4 weeks after TPTX. Thirteen endocrine organs from mice with recovered serum PTH were examined. The thymus from control mice expressed measurable and detectable Pth transcripts; however, the Pth transcript level was substantially elevated in tissue from TPTX mice. Western blot analysis of the thymus demonstrated a reproducible and distinct PTH band in thymus tissue that was significantly increased after TPTX. To directly confirm the identity of the distinct PTH band, immunoprecipitated proteins were isolated and subjected to tandem mass spectrometry. After fragmentation and direct peptide sequencing, PTH peptides PTH(1-13) and PTH(54-70), diagnostic for PTH, were identified. These data demonstrate that the murine thymus produces PTH and that after TPTX the thymus becomes the major source of serum PTH, compensating for the loss of the parathyroid glands and returning circulating PTH levels to normal. Multiple myeloma (MM) is the second most common hematological malignancy and is characterized by the proliferation of malignant plasma cells in the bone marrow, the presence of a monoclonal serum immunoglobulin, and osteolytic lesions (1, 2). Approximately 80% to 90% of patients with MM develop bone lesions that cause bone pain, pathologic fractures, spinal cord compression, and hypercalcemia (1, 2). Such lesions result from dysregulated bone remodeling whereby significant increases in osteolytic activity and impaired bone formation, the results of interactions between MM cells and the bone marrow microenvironment, lead to significant bone destruction (2, 3). Accumulated data from preclinical and clinical studies show that proteasome inhibitors exhibit significant anti-MM activity associated with improved bone remodeling (4), driving the idea that proteasome inhibition is an effective strategy for improving bone remodeling in patients with MM. Whereas the proteasome inhibitor bortezomib inhibits myeloma progression in the myeloma severe combined immunodeficiency (SCID)–hu model, bortezomib increases bone mineral density in both myeloma-bearing SCID-hu mice and non–myeloma-bearing wild-type mice (5). Clinically, during the first cycle of bortezomib treatment in people with myeloma, the bortezomib responder group shows serial serum parathyroid hormone (PTH) spikes, whereas PTH in the bortezomib nonresponder group appears unchanged (6). In support of the idea that PTH responses are important for the antimyeloma effect of proteasome inhibition, we demonstrated that treatment with PTH(1-34) significantly suppressed myeloma cell growth, whereas the parathyroid hormone receptor (PTHR)1 antagonist PTH(7-34) significantly abrogated the suppression of myeloma proliferation in vitro (7). In addition, significant PTH(1-34)–dependent inhibition of myeloma progression was observed in myeloma-bearing C57BL/KaLwRij mice (7). It is well known that PTH plays a pivotal role in the regulation of serum phosphate and calcium levels (8). Calcium is essential for a variety of physiological events, and in mammals, serum calcium levels are maintained within a very narrow range (1 to 1.3 mM) (9). Disorders of PTH production result in serious physiological consequences (10, 11). Serum PTH increases the serum calcium concentration by stimulating renal calcium reabsorption and intestinal calcium absorption and by enhancing calcium release from bone. The parathyroid glands were considered the only tissue capable of producing and secreting PTH until Günther et al. (12) generated the glial cells missing 2–deficient mouse (GCM2), a mouse homolog of Drosophila Gcm–deficient mouse. Although Gcm2-deficient mice lack the parathyroid organ, a substantial number of mice survive and grow normally (12). Indeed, these authors reported that surviving mice had detectable and measurable levels of serum PTH along with an expected and mild hypoparathyroidism (12). It was discovered that the thymus of Gcm2-deficient mice expressed Pth transcripts, and the authors concluded that the thymus was the source of circulating PTH in this mouse (12). Subsequently, Liu et al. (13) generated a series of Gcm2-deficient mice in a variety of genetic backgrounds by F1 backcrossing and demonstrated the lethality of the Gcm2−/− mutation in the C57BL/J background. However, backcrossing into the 129S6 background progressively and significantly reduced lethality (13). The conclusion from these authors was that Gcm2-deficient mice survived due to differences in the genetic background, although serum PTH was not detectable in surviving mice (13). These contradictory findings suggest that survival in the absence of parathyroid glands is achievable, yet the explanation of thymic PTH is inconsistent. Given the importance of the PTHR1 axis and the efficacy of PTH(1-34) treatment of inhibiting myeloma progression (7), we tested the effects of thyroparathyroidectomy (TPTX) in the myeloma-bearing C57BL/KaLwRij mouse model. Here, we demonstrate that the murine thymus expresses Pth at a low basal level that is significantly increased by the surgical removal of the parathyroid glands. The amino acid sequence of thymic PTH is identical to parathyroid-derived PTH, and thymic PTH appears responsible for the recovery of circulating PTH in TPTX animals. These studies may provide insight to the role of the PTH-PTHR1 axis in the control of multiple myeloma. Materials and Methods C57BL/KaLwRij mouse and TPTX procedures C57BL/KaLwRij mice were housed at the University of Arkansas for Medical Sciences (UAMS) Animal Facility and bred in house. All animal procedures were reviewed and approved by the UAMS Institutional Animal Care and Use Committee. TPTX was performed in 8- to 12-week-old male and female mice as follows. The mice were anesthetized using 2% to ∼3% isoflurane and placed in a surgical bed, and a midline incision was made. The salivary glands and sternothyroideus muscle were separated at the midline and retracted. The thyroid gland and parathyroid glands, located as a single pair lateral or posterior to the thyroid or on the lateral edge of the thyroid, were excised. The incision was closed with a wound clip, and the mice fasted overnight with deionized water ad libitum (14). Surgical success was assessed by postsurgical measurement of serum PTH (Immutopics Inc., Athens, OH). Tissue harvest/RNA isolation and quantitative real-time polymerase chain reaction At the time of harvest, the heart, lung, liver, spleen, kidney, pancreas, testis, ovary, hypothalamus, pituitary gland, thymus, and adrenal glands were removed and immediately homogenized in TRI Reagent (Molecular Research Center, Cincinnati, OH). Bone marrow cells were harvested from the femur and tibia with a mortar and pestle in ice-cold phosphate-buffered saline (PBS). The crushed bones were rinsed twice with ice-cold PBS, and the cell suspensions were filtered through nylon mesh (70 μm) to remove debris and connective tissue. The filtered and harvested bone marrow cells were centrifuged at 300 g and resuspended in 1 mL TRI reagent. Total RNA was extracted using the TRI reagent protocol according to the manufacturer’s instructions. The integrity of the harvested total RNA was estimated with gel electrophoresis (0.8% agarose gel). The RNA concentration was determined using a Nanodrop spectrophotometer (Thermo Fisher Scientific, Waltham, MA). Complementary DNA was synthesized with SuperScript III reverse transcription (Invitrogen, Carlsbad, CA) according to the manufacturer’s instructions. Quantitative real-time polymerase chain reaction (qRT-PCR) was performed with the Power SYBR Green PCR Master Mix on the Step-One-Plus system (Thermo Fisher Scientific). Custom specific primers (sense: 5′-TACAGCATCAGTTTGTGCATCCC-3′; antisense: 5′-CAGGTGTTTGCCCAGGTTGTG-3′) were used to amplify the murine Pth transcripts, and commercial probe/primer sets were used for 18S ribosomal RNA (rRNA; catalog no. 4318839-1305059; Thermo Fisher Scientific) as an endogenous internal control. The qRT-PCR was performed using standard protocols. The data were normalized to the 18S rRNA, and fold changes in messenger RNA levels were calculated using the 2−ΔΔCt method (15). Immunoprecipitation and Western blot analyses To isolate proteins, the harvested parathyroid glands, thymus, and livers were washed twice with ice-cold PBS and homogenized in 1.5 mL of Tissue Lysis Buffer (T-PER; Thermo Fisher Scientific) containing 1X Protease Inhibitor Cocktail (Roche, Bradford, CT). Tissue lysates were incubated with 1 μg goat anti-mouse PTH antibodies, including anti-mouse PTH(1-12) [catalog no. 20-2320; Research Resource Identifier: AB_2721076; Quidel, San Diego, CA] and anti-mouse PTH(53-84) (catalog no. 20-2310; Research Resource Identifier: AB_2721077; Quidel) or with 1 μg goat immunoglobulin G (IgG; catalog no. ab37373; Abcam, Cambridge, MA) as an isotype control for 30 minutes at 4°C on a rotary shaker. Protein A-Sepharose (catalog no. 101041; Invitrogen) was added, and the samples were incubated with rotation for 45 minutes at 4°C. The complexes were washed several times with PBS and resuspended in 80 μL of gel sample loading buffer [62.5 mM Tris-HCL (pH 6.8), 2% sodium dodecyl sulfate (SDS), 10% glycerol, 200 mM β-mercaptoethanol, and 0.02% bromophenol blue]. Forty microliters of this sample was subjected to SDS–polyacrylamide gel electrophoresis (PAGE) (15% polyacrylamide gel) and transferred to a polyvinylidene fluoride membrane with a Pierce G2 Fast Blotter (Thermo Fisher Scientific). After nonspecific blocking with 5% nonfat milk for 1 hour at room temperature, the membrane was incubated with primary antibodies [goat anti-mouse PTH(1-12) and PTH(53-84), 1:1000; Quidel] in 1% milk overnight at 4°C. The next day, the membrane was incubated with an horseradish peroxidase–conjugated secondary antibody (1:2000; catalog no. PI9500; Vector Laboratories, Burlingame, CA) and visualized by Immobilon Western (Millipore, Billerica, MA) using the C-DiGit Blot Scanner (LI-COR Biosciences, Lincoln, NE). Measurement of serum calcium Blood (100 μL) was collected via the retro-orbital vein and treated with heparin before surgery and at 3, 24, and 72 hours after TPTX. Plasma was isolated by centrifugation at 4000 g for 5 minutes and stored at −80°C until use; plasma stored at −80°C was not thawed more than once. To have sufficient volume sample, 8 μL of plasma was used for assay using the Calcium Colorimetric Assay kit (Sigma Aldrich, St. Louis, MO) according to the manufacturer’s instructions. Enzyme-linked immunosorbent assay for serum PTH Blood (100 μL) was collected via the retro-orbital vein and treated with heparin on days 1, 3, 7, 14, 21, 28, 35, 42, and 49 after TPTX. Plasma was isolated by centrifugation at 4000 g for 5 minutes and stored at −80°C until use; plasma stored at −80°C was not thawed more than once. To have sufficient volume sample, plasma was diluted to 1 volume of PBS. Diluted plasma (20 μL) was used for assay using the Mouse PTH1-84 enzyme-linked immunosorbent assay (ELISA) kit (Quidel) according to the manufacturer’s instructions. The detection limit was determined by average values from dilution buffer alone on multiple independent experiments (n > 5 sets). Tissue preparation and immunohistochemistry for PTH At 49 days after TPTX, when PTH levels returned to the pre-TPTX level, TPTX and control animals were euthanized, and the thymus glands were removed. Each thymus gland was extensively washed with PBS to remove all blood and fixed with 4% paraformaldehyde in 0.1 M sodium phosphate buffer for 1 day at 4°C. After washing with 1× PBS, tissues were dehydrated with ethanol and embedded in paraffin. Paraffin-embedded tissue blocks were sectioned at 6 µm with an RM2155 microtome (Thermo Fisher Scientific). After deparaffinizing, thymus tissue sections were incubated with anti–PTH(1-12) (1:500) antibodies overnight at 4°C, followed by blocking with 1X PBS containing 1% bovine serum albumin and 5% normal donkey serum for 1 hour at room temperature. After washing with 1X PBS, the sections were incubated with biotin-conjugated anti-rabbit IgG (Jackson ImmunoResearch, West Grove, PA) followed by incubation with ABC solutions (Vector Laboratories, Burlingame, CA). The immunoreactivity of antibodies was visualized with a 3,3′-diaminobenzidine substrate kit (Vector Laboratories) according to the manufacturer’s instructions and then counterstained with methyl green (Vector Laboratories). Images were acquired on a Zeiss Axioplan microscope equipped with an AxioCam digital camera and imaging system (Carl Zeiss Microscopy, Thornwood, NY). Mass spectrometry After immunoprecipitation, the proteins were separated with a 4% to 12% Bis-Tris gel. The entire gel lane was excised and cut into 24 slices, and each slice was subjected to in-gel digestion composed of four steps: destaining, reduction, alkylation, and trypsin digestion (16, 17). The resulting peptides were identified with nano–high-performance liquid chromatography (LC) tandem mass spectrometry (MS/MS) using a nanoAcquity UPLC (Waters, Milford, MA) coupled to a Fusion Oribtrap (Thermo Fisher Scientific) mass spectrometer. An integrated spray tip and nano–high-performance LC column (0.075 × 200 mm) (New Objective, Woburn, MA) was packed with 4 μm 90A Jupiter Proteo resin (Phenomenex, Torrence, CA). A 1-cm loading column allowed for 20 μL of sample to load at 4 µL/min followed by peptide separation at 300 nL/min. Peptides were separated over a multislope gradient consisting of 1% to 35% B for 45 minutes and to 90% B for 5 minutes, where buffer A was 0.1% formic acid and buffer B was 99.9% acetonitrile with 0.1% formic acid. Peptides were ionized using a spray voltage of 2.3 kV. The data were acquired in a data-dependent, top speed manner with a full mass spectroscopy (MS) scan at high resolution (500,000) over a mass range of 375 to 1450 in the orbitrap every 2 seconds, interlaced with as many fragment ion scans in the ion trap as time allowed between full MS scans. High-energy collisional dissociation was used for fragmentation at a normalized collision energy of 28 or 31. Replicate analyses were performed, and the number of gel slices analyzed was limited to the molecular weight region of interest (below 50 kDa). After LC/MS/MS analysis, all MS/MS data were processed using MaxQuant software (v220.127.116.11; http://www.coxdocs.org/doku.php?id=maxquant:common:download_and_installation#download_and_installation_guide). The search parameters were set to allow a false discovery rate (FDR) of 1% at the peptide and protein level with the following parameters: precursor ion tolerance of 4 ppm, fragment ion tolerance of 0.50 Da, fixed modifications of carbamidomethyl on cysteine, variable modifications of oxidation on methionine, and up to three missed cleavages of trypsin. The data were searched against a common contaminant database followed by a search against the UniprotKB database for Mouse. The mouse PTH sequence was identified from multiple gel fragments (NCBI Accession: NP_065648). Statistical analysis All statistical analyses comparing two groups were conducted with a Student t test. A P value of <0.05 was considered significant. Prism4 (GraphPad Software Inc., La Jolla, CA) was used for all statistical analyses. Results Serum PTH levels returned to normal after TPTX To monitor changes in serum PTH after TPTX, we performed TPTX on C57BL/KaLwRij mice (8 to 12 weeks old; n = 15). After surgery, drinking water was supplemented for 1 week with 1 M CaCl2 to prevent hypocalcemia. Five mice died due to surgical complications. In TPTX mice (n = 10), serum was collected 3 days before surgery and after surgery on days 1, 3, 7, 14, 21, and 35, and serum PTH was measured with a mouse PTH ELISA kit (Fig. 1A). Serum PTH dropped below the assay’s limit of detection at day 1 and remained low for 3 days. Any animal that did not show the initial drop in PTH to below the limit of detection was removed from the study because this indicated that TPTX was not complete. Within 1 to 2 weeks in the surviving animals, serum PTH level progressively increased to ∼80% of baseline (i.e., presurgery) levels. Serum calcium levels were also measured before and after TPTX (Fig. 1B). The majority of TPTX mice demonstrated significant initial reduction in serum calcium levels (∼77% of presurgery levels 24 hours after TPTX). However, these expected decreases in serum calcium progressively recovered to ∼85% of presurgery levels 72 hours after TPTX. In addition, serum PTH increased in some mice to levels above the presurgery level at week 5. Given that successful TPTX was demonstrated by the rapid decrease in serum PTH and hypocalcemia, the recovery of serum PTH after surgery raised the possibility of an alternative PTH source in the absence of a functional parathyroid gland. Figure 1. View largeDownload slide Serum PTH levels recover after TPTX. (A) Blood (100 μL) was collected from 10 mice 3 days before surgery (0) and on days 1, 3, 7, 14, 21, 35, and 49 after TPTX via the retro-orbital vein and treated with heparin. Plasma was obtained and kept at −80°C until assay. Mouse PTH in the serum was measured with the mouse PTH(1-84) ELISA Kit according to the manufacturer’s protocol (65 pg/mL PTH is the limit of detection). (B) Blood was collected from 24 mice before TPTX and at 3, 24, and 72 hours after TPTX. Serum calcium levels were measured by calcium colorimetric assay according to the manufacturer’s protocol. Student t test was performed for each group comparison against the presurgery group. **P < 0.01; ***P < 0.001. Figure 1. View largeDownload slide Serum PTH levels recover after TPTX. (A) Blood (100 μL) was collected from 10 mice 3 days before surgery (0) and on days 1, 3, 7, 14, 21, 35, and 49 after TPTX via the retro-orbital vein and treated with heparin. Plasma was obtained and kept at −80°C until assay. Mouse PTH in the serum was measured with the mouse PTH(1-84) ELISA Kit according to the manufacturer’s protocol (65 pg/mL PTH is the limit of detection). (B) Blood was collected from 24 mice before TPTX and at 3, 24, and 72 hours after TPTX. Serum calcium levels were measured by calcium colorimetric assay according to the manufacturer’s protocol. Student t test was performed for each group comparison against the presurgery group. **P < 0.01; ***P < 0.001. Pth transcripts were present in the thymus, and Pth expression increased in the absence of the parathyroid gland To determine which tissues were responsible for the observed PTH recovery in TPTX mice, we examined the primary and secondary endocrine organs. Pth gene expression was examined in 12 TPTX mice that demonstrated serum PTH loss and subsequent recovery. The heart, lung, liver, spleen, kidney, pancreas, ovary/testes, thymus, pituitary gland, hypothalamus, adrenal glands, and bone marrow were harvested. Total RNA was extracted, and Pth transcripts were measured with qRT-PCR. Because commercially available Pth primers span exons 2 and 3 with introns of ∼100 bases, it is difficult to differentiate low-frequency Pth transcripts from genomic DNA contamination. To improve the selectivity and sensitivity of qRT-PCR, we designed custom primers that spanned exons 1 and 3 (Fig. 2A). Of the 11 tissues analyzed, Pth transcripts were detected only in the thymus (Fig. 2B). In addition, detectable Pth transcripts were observed in control (non-TPTX) mice. The detectable levels of Pth transcript present in control mice appeared significantly elevated (∼3 times) in mice lacking parathyroid glands and whose serum PTH recovered (P = 0.0074). The validity of the assay was confirmed by comparing thymic Pth transcript levels using a commercial probe (Assay ID Mm00451600_g1; Applied Biosciences) (Fig. 2B). Furthermore, the amplicon from the thymus was sequenced and confirmed as a 100% match to the reported mouse parathyroid gland Pth transcript sequence (data not shown). Figure 2. View largeDownload slide The thymus produces a significant number of Pth transcripts that increase after TPTX. (A) The thymus produced Pth transcripts. Mice in which serum PTH levels recovered (n = 10) were euthanized, and the heart, lung, liver, spleen, kidney, pancreas, testis or ovary, hypothalamus, pituitary gland, thymus, and adrenal glands were harvested. RNA qRT-PCR was performed on the tissue samples using SYBR Green/Taqman Master Mix with custom primers for the mouse Pth transcript and a commercial probe-set for 18S rRNA (endogenous control). Expression data were normalized to 18S rRNA, and fold changes of expression against arbitrary numbers were calculated using the 2−ΔΔCt method (15). (B) Pth transcripts increased after TPTX. Thymi from unoperated mice (control) were harvested, and the same procedures were performed as in (A). qRT-PCR was performed as described with a commercial probe (gray) and custom probes (black). Student t test was performed. **P < 0.01. G, gland. Figure 2. View largeDownload slide The thymus produces a significant number of Pth transcripts that increase after TPTX. (A) The thymus produced Pth transcripts. Mice in which serum PTH levels recovered (n = 10) were euthanized, and the heart, lung, liver, spleen, kidney, pancreas, testis or ovary, hypothalamus, pituitary gland, thymus, and adrenal glands were harvested. RNA qRT-PCR was performed on the tissue samples using SYBR Green/Taqman Master Mix with custom primers for the mouse Pth transcript and a commercial probe-set for 18S rRNA (endogenous control). Expression data were normalized to 18S rRNA, and fold changes of expression against arbitrary numbers were calculated using the 2−ΔΔCt method (15). (B) Pth transcripts increased after TPTX. Thymi from unoperated mice (control) were harvested, and the same procedures were performed as in (A). qRT-PCR was performed as described with a commercial probe (gray) and custom probes (black). Student t test was performed. **P < 0.01. G, gland. PTH proteins were detected in the thymus, and their abundance increased when the parathyroid was removed The presence of Pth transcripts in the thymus of Gcm2-deficient mice has been reported (12, 13). However, no evidence exists to show expression of PTH protein in the thymus. It is difficult to detect PTH protein from the thymus, likely because (1) the amount of PTH protein made in the thymus is much lower than in the parathyroid glands; (2) when the parathyroid gland is absent, PTH protein is not stored but is actively secreted into the circulation; and (3) the commercially available anti-PTH antibody is raised from rat PTH and may not be sensitive enough to detect PTH in the murine thymus. Thus, to detect PTH in the thymus, we acquired an antibody raised against mouse PTH (Quidel) and demonstrated a significant enhancement of sensitivity against mouse serum PTH with ELISA. Western blot analysis with this antibody was performed on thymus tissue extracts from TPTX mice after serum PTH recovered, and we observed a faint but specific band (data not shown). After extensively washing the thymus tissue to remove all serum PTH, the PTH in the thymus was concentrated by immunoprecipitation with the mouse PTH antibody, followed by Western blot. A specific PTH band was observed at the approximate size of PTH from the parathyroid+thyroid extracts and from the immunoprecipitated sample (Fig. 3A). This PTH band was absent from the immunoprecipitated liver sample and from the antibody isotype controls. In agreement with Pth transcript data, TPTX significantly increased the amount of PTH protein (Fig. 3B). Figure 3. View largeDownload slide Thymus-derived PTH increases after TPTX. After serum PTH returned to a pre-TPTX level, TPTX and control mice were euthanized, and parathyroid and thyroid (Thyroid+Parathyroid), liver, and thymus tissues were removed. Harvested tissues were washed extensively in PBS and homogenized in T-PER buffer, and proteins were immunoprecipitated with goat anti-mouse PTH antibodies, including anti-mouse PTH(1-12) and anti-mouse PTH(53-84) or goat IgG as an isotype control. (A) PTH was detected in whole cell lysates (WCL) of the parathyroid+thyroid, after immunoprecipitation from the parathyroid+thyroid, and in the thymus (indicated by arrows). PTH protein was not detected in the liver. (B) Thymi from TPTX mice contained more PTH than thymi from nonoperated (control) mice. (C) Immunohistochemistry showing increased PTH protein production in the thymus after TPTX. Thymus sections were prepared and incubated with anti–PTH(1-12) antibody. The antibody was visualized with 3,3′-diaminobenzidine and counterstained with methyl green. The representative PTH signals are indicated by yellow arrows. (D) PTH-positive cells were enumerated in at least four sections from each group and plotted as mean ± standard error. Student t test was performed. ***P < 0.001. IP, immunoprecipitated; Iso, isotype control antibody; PT, parathyroid glands; PTH*, parathyroid hormone antibody. Figure 3. View largeDownload slide Thymus-derived PTH increases after TPTX. After serum PTH returned to a pre-TPTX level, TPTX and control mice were euthanized, and parathyroid and thyroid (Thyroid+Parathyroid), liver, and thymus tissues were removed. Harvested tissues were washed extensively in PBS and homogenized in T-PER buffer, and proteins were immunoprecipitated with goat anti-mouse PTH antibodies, including anti-mouse PTH(1-12) and anti-mouse PTH(53-84) or goat IgG as an isotype control. (A) PTH was detected in whole cell lysates (WCL) of the parathyroid+thyroid, after immunoprecipitation from the parathyroid+thyroid, and in the thymus (indicated by arrows). PTH protein was not detected in the liver. (B) Thymi from TPTX mice contained more PTH than thymi from nonoperated (control) mice. (C) Immunohistochemistry showing increased PTH protein production in the thymus after TPTX. Thymus sections were prepared and incubated with anti–PTH(1-12) antibody. The antibody was visualized with 3,3′-diaminobenzidine and counterstained with methyl green. The representative PTH signals are indicated by yellow arrows. (D) PTH-positive cells were enumerated in at least four sections from each group and plotted as mean ± standard error. Student t test was performed. ***P < 0.001. IP, immunoprecipitated; Iso, isotype control antibody; PT, parathyroid glands; PTH*, parathyroid hormone antibody. To determine which cells produce thymic PTH, thymi from control and TPTX mice were subjected to immunohistochemistry with the same anti-PTH antibody. PTH was observed in the medullary thymic epithelial cells as previously reported (13) (Fig. 3C). Significantly more PTH-positive cells (48.3 ± 1.2) were present in the thymus of TPTX mice compared with non-TPTX (control) mice (12.6 ± 0.7; P < 0.0001) (Fig. 3D). Thymic PTH was confirmed by MS To validate that the thymus produced bona fide PTH, immunoprecipitated thymic PTH was analyzed by MS/MS. After extensive tissue washing, total protein was immunoprecipitated from thyroid+parathyroid and whole thymus extracts with the anti-PTH antibody. The immunoprecipitated samples were separated by SDS-PAGE, and peptides were generated via in-gel trypsin digestion. The resulting pool of tryptic peptides was subjected to LC/MS/MS using a Fusion Orbitrap mass spectrometer. The raw MS data were processed using the MaxQuant (v18.104.22.168) software package (18, 19), allowing an FDR of 1% at both the peptide and protein levels. In both tissue samples, several PTH peptides were identified (Fig. 4). In the thyroid+parathyroid sample, three specific PTH fragments were identified: PTH(1-20), PTH(26-44), and PTH(54-82). In the thymus, two PTH peptides were identified: PTH(1-13) and PTH(54-80). Although the data were searched with an FDR threshold of 0.01, both peptides were identified with much higher confidence, having posterior error probabilities (18, 19) of 2e-4 and 8e-5, respectively. In addition, the measured m/z of the peptides was <1 ppm from the predicted values, further validating the identification (data not shown). These data confirm the presence of PTH in the thymus that was identical to parathyroid-derived PTH. Our observation of more PTH peptides in the thyroid+parathyroid sample is reflective of the higher abundance of PTH in that tissue. Figure 4. View largeDownload slide PTH isolated from the thymus was identified by MS. Immunoprecipitated samples from thyroid and parathyroid (Thyroid+PT) glands and the thymus were separated by SDS-PAGE, and peptides were generated with in-gel digestion. The pool of resulting peptides was subjected to LC/MS/MS using a Fusion Orbitrap mass spectrometer. The raw MS data were processed using the MaxQuant (v22.214.171.124) software package (18, 19), allowing for an FDR of 1% at both the peptide and protein levels. In the Thyroid+PT sample, three specific PTH fragments were detected: PTH(1-20), PTH(26-44), and PTH(54-82). In the thymus, two PTH peptides were detected: PTH(1-13) and PTH(54-80). Detected fragments are shown in blue; the predicted signal peptide and predicted PTH peptides are shown in red and white, respectively. Figure 4. View largeDownload slide PTH isolated from the thymus was identified by MS. Immunoprecipitated samples from thyroid and parathyroid (Thyroid+PT) glands and the thymus were separated by SDS-PAGE, and peptides were generated with in-gel digestion. The pool of resulting peptides was subjected to LC/MS/MS using a Fusion Orbitrap mass spectrometer. The raw MS data were processed using the MaxQuant (v126.96.36.199) software package (18, 19), allowing for an FDR of 1% at both the peptide and protein levels. In the Thyroid+PT sample, three specific PTH fragments were detected: PTH(1-20), PTH(26-44), and PTH(54-82). In the thymus, two PTH peptides were detected: PTH(1-13) and PTH(54-80). Detected fragments are shown in blue; the predicted signal peptide and predicted PTH peptides are shown in red and white, respectively. Discussion Previously, we and others demonstrated that proteasome inhibition suppresses myeloma progression clinically (2, 4, 6, 20–23). In addition, treatment with the proteasome inhibitor bortezomib increases bone mineral density in the myeloma SCID-hu mouse model and in nonmyeloma-bearing mice (5). During the first cycle of bortezomib treatment in patients with MM, serial serum PTH elevations were seen in one bortezomib-responder patient group (6), suggesting a link between bortezomib and PTH. In support of this idea, PTH has recently been shown to play an important role in the antimyeloma activity of proteasome inhibitors by suppressing myeloma cell growth (7). Because 5TGM1-transplanted C57BL/KaLwRij mice display many symptoms of human myeloma, including the devastating bone lesions (7, 24–26), the role of PTH in myeloma development was evaluated in vivo by removing the parathyroid glands, which are the primary source of endogenous PTH. Surprisingly, TPTX significantly suppressed myeloma progression in 5TGM1 myeloma-bearing mice (27). After TPTX, a significant portion of the mice survived the dramatic drop in serum PTH. Indeed, many mice had PTH levels return to ∼80% of the presurgery levels within 2 to 3 weeks after TPTX. After total TPTX, serum PTH levels dropped below the assay detection limit (65 pg/mL), providing confidence that the parathyroid glands had been removed. Although serum PTH returned to ∼80% of baseline (determined 3 days before surgery) in most animals within 1 to 2 weeks, it continued to increase in some animals beyond their original baseline for up to 5 weeks after surgery. The recovery of PTH levels after TPTX suggests that serum PTH originates from tissue(s) other than the parathyroid glands. To test the hypothesis of an extraparathyroid source of serum PTH, Pth transcripts were measured by qRT-PCR in samples from various endocrine organs after TPTX. The commercially available probes could not reliably detect Pth transcripts either because the probes lacked sensitivity to detect Pth transcripts from other tissue(s) or because they were unable to differentiate Pth transcripts from genomic DNA contamination. Therefore, using specific primers that span exons 1 and 2, assay sensitivity was improved. Because intron 1 of the murine Pth gene is >1 kb in length (28), genomic Pth DNA contamination is not amplified during conventional qRT-PCR. With this custom primer set, Pth transcripts were detected in thymic tissue. The surgical removal of thyroid and parathyroid tissues allowed comparison of the thymus of unoperated (control) and TPTX mice. Thymus tissue from control animals also expressed PTH, although at a significantly lower level than TPTX animals. Pth transcripts were not detected in any of the other tissues tested. Thymic PTH was initially reported in Gcm2-deficient mice, which are developmentally devoid of functional parathyroid glands (12). In this mouse, Pth transcripts were identified in the thymus, and circulating serum PTH was detected (12). However, another study reported the presence of Pth transcripts in the thymus but did not detect PTH in the serum (13). These authors conclude that there is no compensation of serum PTH from the thymus in Gcm2-deficient mice (13). The discrepancies over serum PTH may arise from the different PTH ELISA assays used. Günther et al. (12) did not report the assay kit used, and Liu et al. (13) used a rodent PTH(1-34) ELISA kit (Quidel). The antibody commonly used to detect mouse PTH is raised against rat PTH rather than mouse PTH. Although there is considerable similarity among the assays (5 amino acid variations among 84), relatively small variations may reduce the sensitivity of this antibody for mouse PTH (29). This difference in sensitivity may not affect detection within the normal range of serum PTH; however, it may be an important criterion for the detection of thymus-derived PTH. Specific anti-mouse PTH antibodies were used for the experiments described herein and reliably and reproducibly detected serum PTH in TPTX mice. To ensure that the protein detected in the thymus was reliably and incontrovertibly identical to parathyroid-derived PTH, mouse PTH amino acid sequences were identified by MS. In the current study, surgery, rather than genetic deletion, was used to remove the parathyroid glands from mice. The surgical approach avoids complications that could arise due to the absence of PTH during development. However, this approach may raise other issues, such as the incomplete ablation of parathyroid glands (although serum PTH and calcium dropped precipitously in our mice). Careful evaluation of Pth transcripts and protein levels in the thymus confirmed that the murine thymus produces PTH under normal conditions and after TPTX. After TPTX, the thymus becomes the major source of PTH, compensating for the loss of parathyroid-derived PTH, and returns circulating PTH levels to near normal. Should such an effect of extra parathyroid-derived PTH exist in humans, it may contribute to the control of myeloma progression and myeloma bone disease. Abbreviations: Abbreviations: ELISA enzyme-linked immunosorbent assay FDR false discovery rate LC liquid chromatography LC/MS/MS liquid chromatography/tandem mass spectrometry MM multiple myeloma MS mass spectroscopy MS/MS tandem mass spectrometry PAGE polyacrylamide gel electrophoresis PBS phosphate-buffered saline PTH parathyroid hormone PTHR parathyroid hormone receptor qRT-PCR quantitative real-time polymerase chain reaction rRNA ribosomal RNA SCID severe combined immunodeficiency SDS sodium dodecyl sulfate TPTX thyroparathyroidectomy UAMS University of Arkansas for Medical Sciences Acknowledgments Financial Support: Thymic PTH was micro-sequenced at the UAMS Sequencing Core Facility, which is supported in part by Translational Research Institute Grant UL1TR000039 through the National Institutes of Health (NIH) National Center for Research Resources and the National Center for Advancing Translational Sciences. L.J.S. was supported by NIH Grant R01CA166060. 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Endocrinology – Oxford University Press
Published: Apr 1, 2018
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