HER2) gene amplification in circulating tumor cells (CTCs) and disseminated tumor cells (DTCs) might be useful for modifying Herceptin therapy in breast cancer. In the process of investigating the utility of a microfluidic platform for detecting HER2 gene amplification in these cells, we observed novel results on discordance of HER2 status. Peripheral blood (8.5 mL) and bone marrow (BM) (7.5–10 mL) were collected prospectively from patients with clinical stages I–IV breast cancer. Mononuclear cells were recovered, stained with cytokeratin (CK), CD45, and DAPI, and processed through microfluidic channels for fluorescence in situ hybridization (FISH). A ratio of HER2:CEP17 >2 in any CK+/CD45 or CK−/CD45 cell was regarded as positive for HER2 gene amplification. Peripheral blood from 95 patients and BM from 78 patients were studied. We found CK+/CD45−/DAPI+ CTCs in 27.3% of patients. We evaluated HER2 gene amplification by FISH in 88 blood and 78 BM specimens and found HER2+ CTCs in 1 of 9 (11.1%) and HER2+ DTCs (27.2%) in 3 of 11 patients with HER2+ primary tumor. Among patients with a HER2− primary tumor, 5 of 79 had HER2+ CTCs (6.3%) and 14 of 67 had HER2+ DTCs (20.8%). The overall rate of discordance in HER2 status was 15% between primary tumor and CTCs and 28.2% between primary tumor and DTCs. HER2 was amplified in CTCs and DTCs in a portion of both HER2+ and HER2− primary tumors. HER2 discordance was more frequent for DTCs. The clinical implications of evaluating HER2 status in CTCs and DTCs in breast cancer needs to be established in prospective clinical trials. The cell enrichment and extraction microfluidic technology provides a sensitive platform for evaluation of HER2 gene amplification in CTCs and DTCs.Human epidermal growth factor receptor 2 ( Mammographic screening and systemic chemotherapy in the adjuvant and neoadjuvant settings have prolonged survival of patients with breast cancer. However, many patients experience disease relapse and progression despite such treatment. Clinical trials have indicated that a cure can be achieved in a proportion of patients by personalizing their treatments. Yet, the tests needed to facilitate personalized treatment in breast cancer, such as quantification and biomarker assessment of minimal residual disease in blood and bone marrow (BM), have not been fully realized.The occurrence of circulating tumor cells (CTCs) in blood and disseminated tumor cells (DTCs) in BM in patients with early or advanced breast cancer is well recognized. These tumor cells most likely play an important role in the complicated process of metastasis Despite early recognition of the potential utility of evaluating CTCs, isolating these cells has remained a challenge because of their extreme rarity in blood. Their estimated frequency is only one cell per 1 billion blood cells It is well recognized that tumor cells undergo epithelial-to-mesenchymal transformation (EMT) as part of the metastatic process BM has long been recognized as an important homing organ for different types of malignant epithelial tumors. The presence of micrometastases in BM at surgery has been shown to be an independent prognostic factor in breast cancer HER2) gene amplification in these cells.The OncoCEE™ (Cell Enrichment and Extraction) microfluidic platform  has been developed and been shown to efficiently capture and detect CTCs Samples of peripheral blood and BM were collected from patients with operable breast cancer in a prospective protocol approved by the institutional review board of The University of Texas MD Anderson Cancer Center . Blood was collected into 8.5-mL vacutainer tubes containing the anticlumping agent CEE-Sure™  and shipped to Biocept's CLIA/CAP–accredited laboratory for processing. BM aspirates (7.5–10 mL) were collected from the bilateral anterior superior iliac crests prior to any surgical manipulation of the primary tumor and were put into tubes containing ethylenediaminetetracetic acid. BM specimens were processed at MD Anderson Cancer Center using a standard density-gradient procedure; a portion of each final cell pellet was shipped to Biocept for DTC capture and analysis.g for 10 min yielded final cell pellets, which were subsequently subjected to tumor cell capture within the OncoCEE™ microchannels. Both CTCs and DTCs captured in this way were then subjected to FISH for detection of HER2 gene amplification.Tumor cells were enriched by density-gradient separation using Percoll (blood) or Ficoll–Hypaque (BM) solution. Following separation, both sample types were incubated with Fc blocker (100 μg/mL) and a 10-antibody capture cocktail  for 30 min at room temperature. The CEE-Sure™ anticlumping reagent was added to the BM samples during the antibody incubation period. Both sample types underwent a wash and centrifugation followed by incubation with biotinylated secondary antibody for 30 min. Three additional washes with phosphate-buffered saline solution/casein buffer and centrifugation at 400 The OncoCEE™ microchannel technology has been described previously For each peripheral blood pellet, captured CTCs were stained with a mixture of pan-CK , anti-CK 7/17 (clone C-46), anti-CK 18 (clone DA/7), and anti-CK 19 (clone A53-B/A2) antibodies labeled with AlexaFluor-488. The CTCs were costained with anti-CD45 labeled with Alexa-594 and then counterstained with DAPI III  for immediate enumeration and localization by microscopic analysis. All CK+/CD45−/DAPI+ cells were classified as CTCs. The precise location (X and Y stage coordinates) of each CTC was recorded, permitting relocalization of cells following FISH for nuclear interrogation. The CTCs were visualized and imaged by an Olympus Bx51 fluorescent microscope  at 200× magnification.For each BM pellet, antibody-labeled DTCs were similarly captured by the streptavidin-coated microchannels but were then immediately processed for FISH. Captured DTCs were not subjected to immunostaining to detect CK and/or CD45. All nuclei were counterstained with DAPI III as part of the FISH procedure (described in the next section).HER2 DNA Probe Kit, which consists of two direct-labeled probes (Abbott Molecular) specific to the centromere of chromosome 17 (CEP17-SpectrumGreen) and the HER2 locus (SpectrumOrange). The FISH analysis for CTCs was performed in two steps. First, each CK+/CD45−/DAPI+ cell was relocated and scored for the number of signals corresponding to each probe. All CK−/CD45−/DAPI+ cells (representing possible CTCs lacking CK and CD45 expression) were then scored. This strategy allowed simultaneous scoring of both CK+/CD45−/DAPI+ and CK−/CD45−/DAPI+ CTC phenotypes for the CEP17 and HER2 probes. Because there are no stain criteria for DTCs, all captured DTCs were subjected to FISH.The microchannels were processed for FISH using the FDA-approved PathVysion HER2:CEP17 ratios were recorded, and all cells whose ratio was more than or equal to 2.0 were regarded as positive for HER2 gene amplification.Both CTCs and DTCs were visualized and imaged by the Olympus Bx51 fluorescent microscope at 400× and 600× magnification. HER2 status of CTCs and DTCs was compared with that of the primary tumor. Discordance in HER2 status was calculated based on the number of patients with HER2+ primary tumors with no evidence of HER2 gene-amplified CTCs or DTCs and those patients with HER2− primary tumor with, however, the presence of HER2 gene-amplified CTCs or DTCs.The Matched samples of peripheral blood and BM from 70 patients, peripheral blood alone from 25 patients, and BM alone from eight patients with operable breast cancer, were utilized in the study. Of the 95 patients whose blood samples were processed and subjected to CTC capture using the OncoCEE™ microfluidic device, 26 (27.3%) had CK+/CD45−/DAPI+ CTCs , in numbHER2 in 88 of the 95 patients from whom blood was collected for the study. In 70 of these 88 patients, matched specimens of BM could be evaluated for HER2 by FISH. The details of the clinicopathologic features of the primary tumor are summarized in We could perform FISH for HER2-amplified CTCs in 6 of the 88 (6.8%) patients from whom peripheral blood could be tested for HER2 gene amplification status by FISH using the OncoCEE™ device. The HER2:CEP17 ratios in the gene-amplified CTCs ranged from 2.0 to 18.3. HER2 gene amplification was observed only in CK−/CD45−/DAPI+ cells in two patients, while it was detected only in CK+/CD45−/DAPI+ in the remaining four patients. Among these six patients with HER2-amplified CTCs, four had their primary tumor staged T1N0 and two as T3N0. The primary tumor was positive for HER2 gene amplification in only one of these patients. Therefore, while HER2+ CTCs were found in 1 of the 9 HER2+ primary tumor, they were encountered in 5 of the 79 HER2− primary tumors. The details of the HER2 gene-amplified CTCs and correlation with HER2 status of the primary tumor is summarized in We observed HER2-amplified FISH signals in all the captured cells. HER2 gene amplification by FISH was observed in 17 of 78 (21.7%) patients for whom a BM specimen was available, with HER2:CEP17 ratios ranging from 2.0 to 15.0. The primary breast tumor was positive for HER2 by FISH in 3 of these 17 (17.6%) patients, and all of these patients were staged as T1N0. In the remaining 14 cases of HER2+ DTCs, the primary tumor was negative for HER2 gene amplification. These 14 tumors were staged as T1N0 (n = 7), T1N1 (n = 2), T2N0 (n = 2), T2N1 (n = 1), T2N3 (n = 1), and T3N0 (n = 1). Therefore, HER2+ DTCs were found in 3 of the 11 patients with HER2+ primary tumors and in 14 of the 67 patients with HER2− primary tumors. The details of the HER2 gene-amplified DTCs and correlation with HER2 status of the primary tumor is summarized in Initially, CK/CD45 staining was attempted on BM specimens, but we observed positive CK staining in cells that were clearly of hematopoietic origin, which indicated frequent false-positive labeling of nonepithelial hematopoietic cells for CK and other antibodies included in the antibody cocktail. Therefore, we could not reliably ascertain the frequency of CK+/CD45−/DAPI+ cells in BM as we could in the peripheral blood. However, we could accurately interpret HER2 status between the primary tumor and CTCs or DTCs included patients in whom the primary tumor was HER2+, but HER2+ CTCs or DTCs were not detected and patients in whom the primary tumor was HER2−, but the CTCs or DTCs were positive for HER2 gene amplification. Overall, 1 of 9 (11.1%) patients with HER2+ primary tumor had HER2+ CTCs and 3 of 11 (27.2%) patients had HER2+ DTCs. Only one patient had concurrent HER2+ CTCs and DTCs. Among patients with a HER2− primary tumor, 5 of 79 (6.3%) had HER2+ CTCs and 14 of 67 (20.8%) had HER2+ DTCs; one patient had concurrent HER2+ CTCs and DTCs. The overall discordance of HER2 gene amplification including patients with HER2+ primary tumor with HER2− CTCs and DTCs and HER2− primary tumor with HER2+ CTCs and DTCs amounted to 15% for CTCs and 28.2% for DTCs.Discordance in the HER2 gene amplification were 15% between primary breast tumor and residual tumor cells in peripheral blood  and 28.2% between primary breast tumor and residual tumor cells in BM . These results confirm that a proportion of patients with breast cancer harbor tumor cells in blood and/or BM, and that the HER2 status of these residual tumor cells can be different from that of the primary tumor.Overall rates of discordance of HER2 status in CTCs and DTCs and their possible role in the personalized treatment of breast cancer is well recognized. Phenotypic characterization of CTCs and DTCs in breast cancer has been reported by previous studies using techniques such as immunofluorescent or immunocytochemical staining to assess HER2 protein overexpression and FISH or polymerase chain reaction (PCR) to evaluate HER2 gene amplification. FISH is considered to be the standard for accurate determination of HER2 status in the selection of patients for trastuzumab therapy because of its low rates of false-negative or false-positive results. Preanalytic and analytic factors yielding inaccurate results can be a major problem with the immunostaining, immunofluorescence, and PCR techniques. Very few studies have reported using FISH for detecting HER2 status of CTCs or DTCs in breast cancer. The current study is one of the largest.The potential utility of evaluating HER2 expression of CTCs and, to a lesser extent, DTCs in breast cancer HER2, MUC1, and GA733-2). Only two small studies have used FISH to evaluate HER2 gene amplification status in CTCs, in both cases following enrichment of EpCAM-expressing cells from peripheral blood by immunomagnetic separation HER2 amplification was regarded as positive for HER2.Several reports have compared HER2 evaluation of CTCs have varied across studies. Studies including patients with locally advanced or metastatic disease reported HER2+ CTCs in 24–47% of the patients, with HER2 status discordance rates ranging from 18% to 58%. In patients with operable breast cancer, 7.9–29% have had HER2+ CTCs, with discordance rates ranging from 3.7% to 44.0%. The 3.7% discordance rate, which corresponded to the 7.9% frequency of HER2+ CTCs, was reported by Ignatiadis et al. HER2 status of CTCs HER2 in primary breast tumors varies from 92% to 98% in the reported studies HER2 could be related to several factors. True biologic differences between RNA levels and DNA gene amplification, analytic variability of two different testing methods, extent of chromosome 17 polysomy, and dilution of mRNA obtained from tumor with those from other nonneoplastic elements can all lead to discordant results The results of HER2 gene amplification in DTCs of patients with operable breast cancer is perhaps the first in the literature. The four previous reports of HER2 expression in DTCs in patients with operable breast cancer utilized immunofluorescence and immunocytochemical staining with a HER2-specific antibody and RT-PCR for HER2 mRNA evaluation HER2 discordance in DTCs in those studies ranged from 38% to 44%. Our 28.2% discordance is lower than these studies, which again is most likely related to the different techniques used for evaluation of the HER2 status. The lower discordance rate found in our study is unlikely due to the representation of patients who had early-stage disease without evidence of metastatic disease because of possible independent modes of dissemination through the hematogenous and lymphatic routes HER2 gene-amplified DTCs in 20.8% of patients, including 25% of those with a HER2+ primary tumor and 20.8% of those with a HER2− primary tumor.Our report of HER2+ CTCs and DTCs in patients with a HER2− primary breast tumor has been reported by all previous studies irrespective of the patient population studied, but the frequency of HER2+ CTCs and DTCs in these patients has varied among these studies. HER2+ CTCs and DTCs were encountered in 19–39% of patients with locally advanced or metastatic disease and in 3.7–31% of patients with operable breast cancer, respectively. The 6% and 21% rates of HER2+ CTCs and DTCs, respectively, that we encountered in patients with a HER2− primary tumor fall within these previously reported ranges.The occurrence of HER2+ CTCs or DTCs in HER2− primary tumors is extensively debated. While the underlying cause of this discordance is not entirely clear, clonal selection of rare HER2+ cells in the primary tumor following treatment is considered to be a possibility in patients with metastatic breast cancer. Genetic instability of the cells constituting minimal residual disease, resulting in acquisition of HER2 in the course of the disease, has been suggested as another possibility. The fact that some of our patients with a HER2− primary tumor demonstrated HER2+ CTCs and DTCs despite having early-stage disease clearly suggests that clonal selection or acquisition due to genetic instability can be encountered very early in the course of the disease. Meng et al., in their study of CTCs detected by FISH in patients with metastatic breast cancer, found lower ratios of HER2:CEP17 in CTCs than in the corresponding primary tumor. Our results clearly indicate that the extent of amplification in CTCs and DTCs can vary relative to that in the primary tumor.The exact cause of HER2+ DTCs to occur more often than HER2+ CTCs in our patient population . HER2+ CTCs and DTCs occurred simultaneously in only two patients, and more commonly in either blood or BM. These results establish the importance of evaluating both CTCs and DTCs when identifying breast cancer patients with discordant HER2 expression. In our study, discordance in HER2 status between HER2− primary breast tumors and minimal residual disease in blood or BM was contributed more often by DTCs than by CTCs (28.2% vs. 15%).Notably, none of the previous studies evaluated CTCs and DTCs simultaneously in patients with breast cancer. We found HER2 gene amplification by FISH. Our identification of CTCs and DTCs with HER2 gene amplification using the OncoCEE™ microchannel system needs further validation in early and advanced breast cancer. This system provides a robust platform for detailed phenotypic and genotypic characterization of intact CTCs and DTCs for accurate evaluation of HER2 status. The clinical utility of this platform for determining HER2 gene amplification by FISH in CK-positive and -negative CTCs and DTCs needs to be tested in large, prospective, multi-institutional clinical trials.The OncoCEE™ microfluidic platform has been shown to be highly efficient and sensitive for the capture of CTCs and now, as shown by our results, for DTCs. The inner surface of the microfluidic channels is coated with streptavidin to which any single biotinylated antibody, or any combination of antibodies, can attach. Mononuclear cells isolated from blood and BM were enriched for tumor cells using an antibody cocktail that included EpCAM. This antibody cocktail helped drive tumor cell capture by increasing cell surface antigen densities. As a result, both epithelial tumor cells expressing low levels of EpCAM and cells with a mesenchymal phenotype lacking EpCAM were captured simultaneously from the same sample. Therefore, we recovered both CK-positive and -negative CTCs and DTCs that were suitable for subsequent testing for


Saccharomyces cerevisiae plays a central role in zinc homeostasis by controlling the expression of genes involved in zinc metabolism. Zap1 is active in zinc-limited cells and repressed in replete cells. At the transcriptional level, Zap1 controls its own expression via positive autoregulation. In addition, Zap1's two activation domains are regulated independently of each other by zinc binding directly to those regions and repressing activation function. In this report, we show that Zap1 DNA binding is also inhibited by zinc. DMS footprinting showed that Zap1 target gene promoter occupancy is regulated with or without transcriptional autoregulation. These results were confirmed using chromatin immunoprecipitation. Zinc regulation of DNA binding activity mapped to the DNA binding domain indicating other parts of Zap1 are unnecessary for this control. Overexpression of Zap1 overrode DNA binding regulation and resulted in constitutive promoter occupancy. Under these conditions of constitutive binding, both the zinc dose response of Zap1 activity and cellular zinc accumulation were altered suggesting the importance of DNA binding control to zinc homeostasis. Thus, our results indicated that zinc regulates Zap1 activity post-translationally via three independent mechanisms, all of which contribute to the overall zinc responsiveness of Zap1.The Zap1 transcription factor of  The activity of transcriptional regulatory proteins can be regulated by a variety of mechanisms. These include the control of transcription factor expression, stability, subcellular localization, DNA binding activity, and activation domain function. While the activity of many transcription factors is regulated by only a single mechanism, some are regulated at multiple levels. Multiple levels of regulating a single transcription factor allow that factor to respond to different signals. For example, the C/EBP transcription factors are regulated at transcriptional, translational, and post-translational levels Another advantage of multiple regulatory mechanisms controlling the activity of a transcription factor is the combined effects each mechanism can have on the response of that factor to a single stimulus. For example, the yeast Pho4 transcription factor is regulated in response to phosphate status by phosphorylation at four different sites in the protein E. coli, zinc homeostasis is accomplished largely through the transcriptional control of zinc uptake and efflux transporters E. coli cells strive to maintain essentially no free zinc in their cytosol In this report, we address the multiple levels of regulation that control the yeast Zap1 protein in response to zinc status. Zinc is an essential nutrient for all organisms because of its many functions as a structural and catalytic cofactor. Zinc is also potentially toxic to cells when accumulated in high amounts. The essential but potentially toxic nature of this metal necessitates precise homeostatic control mechanisms. In Saccharomyces cerevisiae. The Zap1 transcriptional activator is a central player in the adaptation of these cells to zinc-limiting conditions. Zap1 activates the expression of as many as 80 genes We know much about zinc homeostasis in eukaryotes from studies of the yeast 2H2 zinc fingers (Znf). Znf3-7 are located at the C-terminus (residues 705–880) and comprise the DNA-binding domain. This domain is responsible for specific recognition of Zinc Response Elements, or ZREs, found in one or more copies in Zap1's target promoters 5′-ACCTTNAAGGT-3′ and we have shown previously that Zap1 binds to this palindromic sequence as a monomer and not as a dimer Zap1 is an 880-residue protein containing seven CZinc-Responsive Domain of AD1 (ZRDAD1) that is required for controlling AD1 function AD2) includes both Znf1 and Znf2. Studies to date indicate that zinc regulates AD1 and AD2 directly by binding to ligand residues in their respective ZRDs to repress activation domain function Zap1's two activation domains, AD1 and AD2, are responsible for increasing gene expression in response to low zinc. AD1 lies between residues 207 and 402 and is embedded within a larger region (residues 182–502) termed the DBD) was zinc regulated While regulation of AD1 and AD2 clearly plays an important role in Zap1 zinc responsiveness, other levels of control may also contribute. For example, at the transcriptional level, Zap1 induces its own expression in low zinc via positive autoregulation ZRT1-lacZ reporter is highly induced by zinc limitation and expressed at decreasing levels as the concentration of zinc in the medium is increased (zap1Δ cells expressing wild-type Zap1 at a low level from the GAL1 promoter (pZap1WT) showed a similar response. Although this promoter is highly induced by galactose, we grew these cells in glucose where the level of Zap1 expression was similar to the level observed in zinc-replete wild-type cells expressing Zap1 from its own promoter zap1Δ cells expressing a mutant form of the protein (Zap1TC) in which AD1 and AD2 function is made constitutive by mutating regulatory residues required for the zinc responsiveness of those domains ZRT1 and ZPS1, two chromosomal targets of Zap1, were assayed under low and high zinc conditions (2+ in the medium available to cells is less than what is found in standard yeast culture media (i.e. YPD or SD) CMD1, encoding calmodulin, was used as a negative control and its levels were not altered by zinc status.In wild-type cells that express Zap1 from its own promoter, a Zap1-regulated TC protein and the GAD-Zap1DBD fusion in vivo dimethyl sulfate (DMS) footprinting to examine occupancy of a Zap1 binding site (ZRE1) in the ZRT1 promoter zap1Δ mutant grown in zinc-limiting  (see The observation that the Zap1ion) see . Less thZAP1 mRNA ZAP1 transcriptional autoregulation does indeed cause corresponding changes in Zap1 protein level. Therefore, the apparent loss of ZRT1 ZRE occupancy observed in wild-type cells grown in high zinc could be due to decreased Zap1 DNA binding activity and/or decreased Zap1 protein level resulting from transcriptional autoregulation.Zap1 controls its own transcription via positive autoregulation. Therefore, the effects of zinc on ZRE occupancy in wild-type cells could result solely from changes in Zap1 protein level. To assess this possibility, we first determined the level of Zap1 protein expressed under these conditions by immunoblot analysis  in zinc finger 4 were mutated to alanines This observation was useful because it allowed us to test the importance of DNA binding regulation to overall Zap1 zinc responsiveness. Zap1 was expressed in a zap1Δ cells expressing either low or high levels of Zap1 were grown in LZM supplemented with 3, 10, 30 and 100 µM Zn and tracer amounts of 65Zn to assess total zinc accumulation. After ∼5 generations of growth in these media, the cells were harvested, washed to remove surface-bound zinc, and assayed for accumulation. As shown in zap1Δ cells expressing low levels of Zap1. These results support the hypothesis that DNA binding control is critical for maintaining zinc homeostasis in these cells.These results show that regulation of Zap1 DNA binding is important for the zinc responsiveness of Zap1 transcriptional regulation. To assess its importance to zinc homeostasis, we assayed zinc accumulation in cells with or without DNA binding control. An effect on zinc accumulation would reflect misregulation of the zinc uptake transporters. Wild-type cells and in vivo DMS footprinting and chromatin immunoprecipitation. One limitation of the footprinting method is that it does not definitively show that the protein protecting the ZRE is Zap1 and not some other transcription factor. However, several observations support the conclusion that the methylation protection observed is due to Zap1 binding. First, no protection was detected in a zap1Δ mutant and protection was constitutive in a Zap1-overexpressing strain. Second, in vitro studies have shown that purified Zap1 binds to ZREs with high sequence specificity and affinity in vivo.In this report, we have shown that Zap1 DNA binding activity is regulated by zinc status. This conclusion is based on our observations that ZRE occupancy is altered in response to zinc without any changes in the total level of Zap1 or the nuclear localization of the protein. ZRE occupancy was assayed using both ZAP1 gene expression is under positive transcriptional autoregulation. Here we confirm that this transcriptional control does indeed alter Zap1 protein levels.With the addition of DNA binding control, we now know that Zap1 activity is regulated by zinc via multiple mechanisms. Two of these mechanisms involve the regulation of Zap1's two activation domains. Our previous studies showed that zinc controls AD1 and AD2 independently of each other. This regulation likely occurs through the binding of zinc directly to residues within and flanking these activation domains. We hypothesize that zinc binding to these domains folds them into inactive conformations that are incapable of recruiting coactivators to Zap1 target promoters. We have also shown in previous studies that ZRT1-lacZ reporter when we eliminated that mechanism of control by expressing Zap1 at a constant low level from the GAL1 promoter. However, we note that the ZRT1 promoter contains four high affinity ZREs and therefore may be less sensitive to changes in Zap1 level than other promoters with lower affinity binding sites. This hypothesis is supported by our recent observation that Zap1 target genes with lower affinity ZREs tend to be induced only under the most severe zinc-limiting conditions when Zap1 protein levels are highest ZAP1 expression may play a significant role on those promoters.Having so many different mechanisms of zinc responsiveness raises the issue of what each of these mechanisms contributes to Zap1 regulation and zinc homeostasis. With regard to transcriptional autoregulation, we found no obvious effect on regulation of a ZRT1 mRNA levels decreased following zinc treatment of zinc-limited cells, suggesting this is not the case .From our analysis of activation domain and DNA binding domain regulation, it is clear that these post-translational mechanisms are integrated to control the overall response of Zap1 to zinc. When comparing AD1 and AD2 function, we have found that AD1 is responsible for dictating the zinc dose response and induction kinetics of most Zap1 target genes in vitro but these experiments did not support the hypothesis. Using both electrophoretic mobility shift assays and surface plasmon resonance analysis, we have determined that Zn2+ at concentrations as high as 10 µM does not inhibit ZRE binding by purified Zap1 . This value is much higher than the nanomolar (or lower) levels of free zinc in vitro but this effect was nonspecific and those levels of zinc also interfered with in vitro DNA binding of control proteins  that function well in zinc-replete cells. Thus, inhibition of DNA binding by zinc ions binding directly to the Zap1 DNA binding domain appears unlikely to be the mechanism that operates in vivo. An alternative model is that DNA binding activity is controlled by post-translational modification, such as phosphorylation. This form of DNA binding regulation has been demonstrated for other C2H2 zinc finger transcription factors, such as Adr1 2H2 zinc finger proteins How might zinc control Zap1 DNA binding? We mapped the regulation of DNA binding activity to the Zap1 DNA binding domain alone. The five zinc fingers of the DNA binding domain are all high affinity structural zinc sites and these domains have metal bound even in zinc-limited cells. Our previous studies demonstrated that binding of zinc by each of the five fingers is required for ZRE binding 2. LZM contains 1 mM EDTA and 20 mM citrate as metal buffers to limit zinc availability. It is important to note that even when supplemented with 1000–3000 µM zinc, LZM has lower free Zn2+ available to cells than is found in standard yeast culture media such as SD or YPD. Regulated expression from the GAL1 promoter in glucose-containing media was achieved using the GEV system. The GEV protein contains the Gal4 DNA binding domain, the VP16 activation domain, and the hormone-response domain of the human estrogen receptor. Treatment of GEV-containing cells with β-estradiol results in increased expression of genes expressed from the GAL1 promoter All yeast strains were grown in either YP medium supplemented with 2% glucose (YPD) or synthetic defined medium with 2% glucose and the appropriate auxotrophic supplements. Limiting zinc medium (LZM) was prepared as previously described ade6 can1 his3 leu2 trp1 ura3), ZHY6 (DY1457 zap1Δ::TRP1) MATa leu2 trp1 ura3 prb1 pep4 prc1 gal2) ZRT1-lacZ reporter construct used in this study was pGI-1 ZRT1 promoter (−521 to the ATG start codon) fused to lacZ. The construction of pZap1WT (previously referred to as pMyc-Zap11–880), a plasmid containing an N-terminal myc-tagged ZAP1 allele under the control of the GAL1 promoter in the vector pYef2, was described previously TC was generated as previously described (21). pZap1Δ17-700 was made by generating the corresponding open reading frame by overlap PCR and inserting the resulting fragment into BstX1-linearized pZap1WT using homologous recombination. To create pPGK-ZRT1, a cloning strategy was used in which the ZRC1 open reading frame in the construct pPGK-ZRC1  was replaced with ZRT1 using homologous recombination. Primers were designed such that the 5′ primer contained 40 bp of homology to the PGK1 promoter region and 20 bp of homology to the 5′ end of ZRT1 and the 3′ primer contained 40 bp of homology to the ZRC1 terminator and 20 bp of homology to the 3′ end of ZRT1. The resulting PCR product was co-transformed into DY1457 with BamHI-digested pPGK-ZRC1 and transformants selected on SD media for the plasmid URA3 marker. Plasmids were then rescued from DY1457 using standard procedures. The resulting construct containing ZRT1 fused to the PGK1 promoter and ZRC1 terminator was confirmed by DNA sequencing.The yeast strains used in this study were DY1457 (MATα 600 = 0.3–0.7) in LZM supplemented with the indicated amount of ZnCl2. Cells were treated with DMS (30 µl/30 ml of culture) for 7 min and then harvested by centrifugation. Isolation of genomic DNA and in vivo footprinting analysis were performed as previously described ZRT1 promoter at positions −481 to −460 relative to the initiation codon. Quantification of band intensities was performed using a Storm 860 PhosphorImager (Molecular Dynamics) with ImageQuant software. Calculation of the fractional protection of residues was performed as follows ZRE/BIstandard)/(BIZRE in zap1Δ/BIstandard in zap1Δ)]×100 where BIZRE and BIstandard refer to the band intensities of the affected ZRE residues and of a control band elsewhere in the primer extension ladder that was unaffected by zinc or ZAP1 genotype. Similar results were obtained when ZRE protection was quantified relative to several different control bands.Thirty ml cultures were grown for 15–20 h to mid-exponential phase (AWT) were grown to an A600 ∼0.5 and then treated with 1% formaldehyde to cross-link protein-DNA complexes. The cross-linking reaction was quenched by adding 125 mM glycine. After two washes with ice-cold PBS, the cells were lysed with glass beads in buffer containing Complete Protease Inhibitor Cocktail (Roche), 1 mM PMSF, and 2 mM benzamidine. Following centrifugation for 10 minutes at 16,000× g, the supernatants were incubated with anti-myc antibody at 4°C overnight and immune complexes isolated with Protein A-Sepharose. The cross-links were reversed in TES and co-immunoprecipitation of specific promoter fragments with myc-Zap1 was assessed by PCR using primers flanking the ZRT1 and ZPS1 ZREs  digestion were resuspended in an 18% Ficoll buffer  and were disrupted using a Dounce homogenizer. Following removal of cell debris by centrifugation at 5000× g for 15 min, nuclei and other organelles were separated from the cytosolic fraction by centrifugation at 25,000× g for 30 min. The pellet was resuspended in 18% Ficoll buffer before being layered onto 35% Ficoll in the same buffer and the nuclei sedimented at 100,000× g for 90 minutes. All buffers during and after the disruption of spheroplasts contained 1 mM PMSF, 1 mM EDTA and the Complete Protease Inhibitor Cocktail (Roche). The protease-deficient strain BJ2168 was required for these experiments due to the extreme sensitivity of Zap1 to proteolytic degradation during sample preparation from wild-type cells. Total protein extracts for immunoblots were prepared by cell disruption and protein precipitation in the presence of trichloroacetic acid as described Protein extracts for immunoblots were prepared by two different procedures. Subcellular fractionation analysis was performed as described previously 32P end-labeled oligonucleotide probe before digestion by S1 nuclease and separation on an 8% polyacrylamide/8 M urea gel. Probes used are listed in RNA was extracted from cells grown to mid-log phase using hot acid phenol extraction. S1 analysis was performed as previously described 600 = 0.3–0.7) in LZM supplemented with the indicated amount of ZnCl2. β-galactosidase activity was measured as described 420×1000)/(min×ml of culture×absorbance of the culture at 595 nm).Cells were grown for 15–20 h to mid-exponential phase , and then counted using a Wallac 1480 Wizard™gamma counter. Cell-associated 65Zn was then adjusted for specific activity and normalized by converting the culture A600 values to cell number with a standard curve.Cells were grown in LZM medium supplemented with the indicated concentration of ZnCl


Lmo2) gene encodes a transcriptional cofactor critical for the development of hematopoietic stem cells. Several distal regulatory elements have been identified upstream of the Lmo2 gene in the human and mouse genomes that are capable of enhancing reporter gene expression in erythroid cells and may be responsible for the high level transcription of Lmo2 in the erythroid lineage. In this study we investigate how these elements regulate transcription of Lmo2 and whether or not they function cooperatively in the endogenous context. Chromosome conformation capture (3C) experiments show that chromatin-chromatin interactions exist between upstream regulatory elements and the Lmo2 promoter in erythroid cells but that these interactions are absent from kidney where Lmo2 is transcribed at twelve fold lower levels. Specifically, long range chromatin-chromatin interactions occur between the Lmo2 proximal promoter and two broad regions, 3–31 and 66–105 kb upstream of Lmo2, which we term the proximal and distal control regions for Lmo2 (pCR and dCR respectively). Each of these regions is bound by several transcription factors suggesting that multiple regulatory elements cooperate in regulating high level transcription of Lmo2 in erythroid cells. Binding of CTCF and cohesin which support chromatin loops at other loci were also found within the dCR and at the Lmo2 proximal promoter. Intergenic transcription occurs throughout the dCR in erythroid cells but not in kidney suggesting a role for these intergenic transcripts in regulating Lmo2, similar to the broad domain of intergenic transcription observed at the human β-globin locus control region. Our data supports a model in which the dCR functions through a chromatin looping mechanism to contact and enhance Lmo2 transcription specifically in erythroid cells. Furthermore, these chromatin loops are supported by the cohesin complex recruited to both CTCF and transcription factor bound regions.The Lim domain only 2 ( Lmo2 gene, have shown that Lmo2 is necessary for embryonic yolk sac erythropoiesis Lmo2 expression is maintained in erythroid cells but down regulated in the T-cell lineage Lmo2 results in the development of T-cell related diseases; indeed Lmo2 is located at a recurrent site of T-cell acute lymphoblastic leukemia (T-ALL) specific translocation Lmo2 when the gene therapy vector integrated near Lmo2Lim domain only 2 (LMO2) is a critical transcriptional regulator of hematopoiesis. Gene targeting experiments conducted to introduce null mutations in the mouse Hbb), α-globin (Hba), retinaldehyde dehydrogenase 2, c-kit and erythroid Kruppel-like factor (Eklf) Previous studies have shown that in erythroid cells LMO2 is usually present as part of a complex with the transcriptional regulators, TAL1, E47, LDB1, and GATA1 Hbb locus control region (LCR) Hbb-b1 gene Hbb LCR is in close proximity to the active Hbb genes (Hbb-b1 and Hbb-b2) whereas the intervening 50 kb of DNA sequence containing the embryonic erythroid cell expressed genes is looped out Hba, Shh, TH2, HoxB1 and olfactory receptor genes Transcription is regulated not only by the sequences immediately upstream of gene transcription start sites (TSS) but in many cases by distal regulatory elements (DRE) which can be located up or downstream of the genes they regulate. In fact genome-wide chromatin immunoprecipitation sequencing (ChIP-Seq) analysis for several transcription factors has revealed that a significant proportion (40–60%) of transcription factor bound regions are located in the intergenic regions of the genome ≥10 kb from a gene TSS Hbb, Igf2/H19 and HoxA, however, CTCF bound regions are generally bound by CTCF in all cell types IGF2-H19 locus and at the developmentally regulated IFNG locus both cohesin and CTCF are required for maintaining higher-order chromatin conformation cis-regulatory elements not occupied by CTCF The chromatin-chromatin looping interactions that regulate cell-type specific gene expression are also present in a cell-type specific manner whereas many of the proteins present at sites of looping interactions are ubiquitously expressed. For example, CTCF participates in intra- and inter-chromosomal looping at individual gene loci including Lmo2 in erythroid cells appears to involve multiple distal regulatory elements. Three alternate upstream promoters have been identified as well as multiple conserved DRE located up to 100 kb upstream of the Lmo2 gene Lmo2 and 1 kb downstream are capable of enhancing reporter gene expression in erythroid cells and in transgenic mice suggesting that strong expression of Lmo2 in hematopoietic cells requires the combined action of upstream DRE and sequences close to the Lmo2 proximal promoter Lmo2 transcription in the context of its endogenous genomic location using the 3C technique. Our results show that multiple upstream DRE interact with the Lmo2 proximal promoter whereas intervening regions are looped away from the proximal promoter. DRE upstream of Lmo2 are bound by multiple transcription factors, p300, and associated with intergenic transcription when Lmo2 is transcribed at high levels in erythroid cells. Interaction between the DRE and the Lmo2 promoter was identified in erythroid cells but not in kidney cells suggesting a link between the looping conformation of the locus and transcriptional regulation of Lmo2. Furthermore, a CTCF and cohesin occupied region upstream of the most distal enhancer (90 DRE) also contacts the Lmo2 proximal promoter region, potentially insulating the neighboring Cell cycle associated protein 1 (Caprin1) gene from interaction with the DRE that enhance Lmo2 transcription.Transcriptional regulation of Lmo2 have been identified in the human genome and confirmed to have enhancer activity in transgenic mice Lmo2 , is located close to the Lmo2 proximal promoter and contains the distal and intermediate promoters as well as the 12 and 25 DRE. The second interaction domain located further upstream of Lmo2, which we term the distal control region (dCR), contains the 70, 75 and 90 DRE as well as a fragment upstream of the 90 DRE. These are similar, though not identical, to clusters I (−90 to −64) and II (−40 to +1) identified by Landry et al. 2009 as enriched in histone H3 acetylation in erythroid cells Next we performed a locus-wide 3C using the ey cells . These fCaprin1 and Lmo2 highlighted the 75 DRE as being bound by multiple transcription factors but did not reveal binding at the 90 and 70 DRE. By contrast our 3C data revealed a broad domain of chromatin-chromatin contacts between the 70, 75 and 90 DRE and the Lmo2 proximal promoter raising questions about the function of this dCR. We did identify additional p300 association at the 70 DRE suggesting that bound transcription factors are present which recruit p300 to this DRE , containing three transcription factor bound DRE , and a proximal interaction cluster (pCR). Furthermore, 3C experiments revealed no significant interactions between the Caprin1 TSS and the DRE suggesting these elements are specific in regulating Lmo2 transcription.Our investigation of chromatin-chromatin interactions throughout the mouse genomic region containing Lmo2 the 75 DRE contained the highest density of transcription factor association as well as p300, RAD21 and a nearby CTCF bound region. Furthermore our 3C data confirmed specific interaction of the 75 DRE with the Lmo2 proximal promoter and interaction of the Lmo2 proximal promoter with the dCR containing the 75 DRE as well as the 90 and 70 DRE. In addition, a CTCF bound region upstream of the 90 DRE is contained in the dCR. Previous studies in circulating erythroid cells of transgenic mice show that the 75 DRE has the strongest enhancer activity in erythroid cells and drives the expression of a reporter gene, cooperatively with the Lmo2 proximal promoter and +1 enhancer element Lmo2 proximal promoter/+1 DRE region to form an erythroid cell specific chromatin loop that includes other regulatory elements and a cluster of CTCF bound regions. This suggests that in the endogenous context elements throughout the dCR coordinately regulate Lmo2 transcription in erythroid cells.Using a combination of ChIP-Seq data for mature erythroid cells and HPC7 hematopoietic progenitor cells we identified multiple transcription factors bound within both the pCR and the dCR. Even considering the HPC7 ChIP-Seq data that showed more transcription factor peaks across the entire region upstream of Lmo2. Interestingly, we did not identify interactions of the 75 DRE with the fragments containing the 25 or 12 DRE. Transgenic analysis by Landry et al. 2009 of these elements did reveal that the 25 and 12 DRE conferred expression in the fetal liver whereas only the 75 DRE conferred expression in fetal liver as well as in circulating blood cells suggesting that these elements have different functional roles in regulating Lmo2 expression. The fact that we did not identify specific interactions between the 75 DRE and the fragment containing the 25 or 12 DRE suggests that the interactions of these two regions with the proximal promoter are mutually exclusive. These mutually exclusive interactions could occur in different sub-populations of cells within the anaemic spleen or the interactions could be dynamic within individual cells with the proximal promoter region alternately contacting the 25–12 region and the 70–90 region similar to the flip-flop of the Hbb LCR between the γ- and β-genes We also identified increased interaction throughout a pCR containing the 25 DRE/distal promoter and 12 DRE/intergenic promoter and the +1DRE/proximal promoter in erythroid cells compared to kidney cells, suggesting that this more proximal region also contributes to transcriptional regulation of endogenous Lmo2 proximal promoter; however the question remains as to which factors are mediating these looping interactions. LMO2 itself is one of the regulatory transcription factors bound to the upstream DRE, specifically at 75, 25 and 12, all of which showed increased interaction with the Lmo2 promoter suggesting an important role for the LMO2 complex in chromatin looping. In support of this, a recent study found LDB1 (a member of the LMO2 complex) at regions of chromatin interaction with the LDB1 bound Hbb-b1 promoter Hbb-b1 and the LCR Lmo2 locus. Cohesin (RAD21) is bound within both the dCR and pCR, specifically at the 75, 25 DRE as well as at the proximal promoter suggesting cohesin bound at the upstream DRE supports erythroid cell specific looping interactions with the Lmo2 proximal promoter.We identified erythroid-cell specific chromatin loops between a dCR and the Lmo2 upstream region, all of which occur within erythroid cell specific interacting domains, though the majority of the CTCF sites were not specific to erythroid cells. This is similar to the findings at the Hbb locus where CTCF bound regions, invariant between cell types, formed cell type specific chromatin loops Caprin1 from interacting with the DRE that enhance Lmo2 transcription in erythroid cells. As overexpression of Caprin1 causes inhibition of cell division it is critical to prevent its aberrant upregulation in rapidly cycling erythroid cells Lmo2 locus being supported by cohesin recruited both to CTCF bound regions  as well as at transcription factor and p300 bound enhancers not associated with CTCF .Cohesin is also recruited to CTCF occupied regions Lmo2 dCR is more reminiscent of the human Hbb LCR Lmo2 dCR may function to facilitate the physical interaction of the dCR with the Lmo2 proximal promoter through recruitment of both regions to a shared transcription factory. This co-localization at a shared transcription compartment would then allow transcription factors recruited to the dCR to influence the basal transcriptional machinery recruited to the Lmo2 proximal promoter thereby enhancing Lmo2 transcription.Our analysis indicates that a large portion of the dCR is transcribed at moderate levels in erythroid cells but not in kidney. Whereas short eRNAs have been identified at enhancers in neuronal cells the broad domain of intergenic transcription we observed throughout the Lmo2-Caprin1 locus adopts a tissue-specific conformation in erythroid cells. This tissue specific organization of the locus brings several DRE into proximity with the Lmo2 proximal promoter while excluding the Caprin1 TSS. A proximal control region, immediately upstream of the Lmo2 proximal promoter, contains the 12 and 25 DRE as well as the distal and intermediate promoters and is more closely associated with the Lmo2 proximal promoter in erythroid cells compared to kidney. Furthermore, a distal control region, covering 39 kb and containing three DRE , forms a strong interaction with the Lmo2 proximal promoter and shares many features with the well characterized Hbb LCR. A CTCF bound region upstream of the 90 DRE flanks the dCR and may function as an insulator preventing the interaction of Caprin1 with the erythroid cell specific Lmo2 enhancers.In conclusion, we found that the mouse All experiments were approved by the University Animal Care Committee (UACC) at the University of Toronto and the Bioscience Local Animal Care Committee (LACC).8) from the spleen of mice 5 days after phenylhydrazine treatment. This treatment induces haemolytic anaemia, as a result of which the spleen becomes the major site of red blood cell production Adult globin expressing mature erythroid cells were isolated from C57/Blk6 mice in large numbers (>1×10Lmo2-Caprin1 locus (RP23-76D2) with the Alpha Aortic Actin 2 BAC clone (RP23-2N15) followed by digestion with HindIII. The digested DNA was then ligated, and purified using phenol/chloroform extraction and ethanol precipitation. HindIII restriction enzyme digestion efficiency was confirmed to be between 85 and 95% efficient at several genomic fragments in anaemic spleen and kidney cells  was subsequently used for quantification. PCR products of the ligated fragments were quantified using real-time quantitative PCR (qPCR) on the Bio-Rad CFX-384 cycler. All data points were generated from an average of three-five independent 3C experiments with the qPCR performed in triplicate. Standard curves were generated by 5 fold serial dilution of the 3C control template and run in parallel with 3C experimental samples. The primers used in qPCR are listed in Gapdh or Epn1 were used for normalization of expression levels. The primers used for real-time RT-qPCR are listed in RNA from anaemic spleen and kidney was isolated using TRIzol, according to the manufacturer`s instructions (Invitrogen). The iScript First strand synthesis cDNA kit from Bio-Rad was used for preparation of random-hexamer primed cDNA. Amplification in qPCR was performed on the Bio-Rad CFX-384 cycler using the standard curve method. The reaction mixture contained 2X Bio-Rad iTaq SYBR green mastermix, 0.3 pM of each primer, 1uL cDNA (10 times diluted from a 20 uL of reverse transcription reaction). The conditions for qPCR were as follows: 94°C for 3 min followed by 40 cycles at 94°C for 30 s, 62°C for 30 s. Expression levels of The 3C data were analyzed by two-way ANOVA using Sigma Plot12. Post tests (Holm-Sidak method) were performed to assess significant differences between anaemic spleen and kidney samples at specific genomic locations.ChIP-Seq raw data for GATA1, KLF1, LDB1, TAL1, and MTGR1 Figure S1The Lmo2/Caprin1 region on mouse chromosome 2. Primers used in chromosome conformation capture (3C) and HindIII restriction sites are shown across the Lmo2/Caprin1 region of mouse chromosome 2. Promoters and distal regulatory elements (DRE) are depicted in red and black respectively. Anchor fragments used in the Caprin1, 75 DRE and Lmo2 3C experiments are marked with an asterisk (*). Distal promoter (pP), proximal promoter (pP).(PDF)Click here for additional data file.Figure S2Distal regulatory elements upstream of Lmo2 overlap transcription factor bound regions in HPC7 hematopoietic progenitor cells. The mouse Lmo2-Caprin1 region. Distal regulatory element (DRE) homology regions are indicated by black boxes joined by a line to delineate the human enhancer construct used in the generation of transgenic mice. Coloured boxes represent peaks identified from transcription factor ChIP-Seq data from HPC7 hematopoietic progenitor cells obtained from Wilson et al. 2010. Proximal promoter (pP), distal promoter (dP).(PDF)Click here for additional data file.Figure S3The Caprin1 TSS does not interact with distal regulatory elements upstream of Lmo2. Quantitative chromosome conformation capture (3C) was performed to detect chromatin-chromatin interactions between the Caprin1 TSS and distal regulatory elements (DRE). The profile of interactions identified in anaemic spleen (red) and kidney (blue) is displayed. Black box indicates the anchor fragment at Caprin1 and alternating intensities of grey boxes indicate the fragments investigated for interactions. Data points are an average of three independent biological replicates. Error bars depict the SEM, no significant differences were identified throughout this region.(PDF)Click here for additional data file.Figure S4CTCF bound upstream of Lmo2 in different cell types. The mouse Lmo2 upstream region on chromosome 2 is depicted with chromosome coordinates shown at the top. HindIII restriction sites are indicated by blue lines. The two Lmo2 promoters are indicated by red boxes. Distal regulatory element (DRE) homology regions are indicated by black boxes joined by a line to delineate the human enhancer construct used in the generation of transgenic mice. Mouse ENCODE ChIP-Seq data from B Ren (Ludwig Inst. for Cancer Research) and M Snyder (Stanford University) for CTCF in different cell types are shown below the DRE. Proximal promoter (pP), distal promoter (dP), murine erythroleukemia cells (MEL differentiated with 2% DMSO), bone marrow (BM), embryonic stem cells (ES-Bruce4), mouse embryonic fibroblasts (MEF).(PDF)Click here for additional data file.Figure S5Restriction digestion efficiency in chromosome conformation capture. Restriction digestion efficiency was between 85 and 95% at several HindIII restriction sites. Lmo2 proximal promoter (pP), Distal regulatory element (DRE), Caprin1 promoter (Caprin1P). “U” denotes a restriction fragment upstream of the indicated element.(PDF)Click here for additional data file.Table S1Coordinates of distal regulatory elements located upstream of the Lmo2 promoter in the mouse genome. Distal regulatory elements are named acording to their distance upstream of the annotated Lmo2 transcription start site overlapping the proximal promoter. Coordinates are given for homology regions identified by BLAT. All fragments were mapped in NCBI m37 mouse assembly (mm9).(PDF)Click here for additional data file.Table S2Coordinates of the Lmo2 proximal and distal promoters in the mouse genome. The coordintes of proximal promoters and distal promoters for the Lmo2 gene in the mouse genome are listed in the table. Coordinates are given for homology regions identified by BLAT. All fragments were mapped in NCBI m37 mouse assembly (mm9).(PDF)Click here for additional data file.Table S3Chromatin immunoprecipitation sequencing data. Transcription factor binding sites have been obtained from three different cell types; differentiated murine erythroleukemia cells (MEL), hematopoietic progenitor cells (HPC7), and GIE-ER4 a GATA1-null erythroblast cell line in which GATA1 activity was restored. CTCF, DNaseI hypersensitivity, p300 and RAD21 data have been obtained from the mouse ENCODE project, sources listed .(PDF)Click here for additional data file.Table S4Relative transcript abundance in intergenic regions upstream of Lmo2.(PDF)Click here for additional data file.Table S5Primers. Specific primers are listed for the chromosome conformation capture (3C) and RT-qPCR analyses. Left primer (L), right primer (R), primers used to test HindIII restriction digestion efficiency are marked as REX.(PDF)Click here for additional data file.


Gnas transcript at the Gnas cluster has been thought to account for the PatDp(dist2) phenotype. But PatDp(dist2) also have two expressed doses of the paternally expressed Gnasxl transcript. Through the use of targeted mutations, we have generated PatDp(dist2) mice predicted to have 1 or 2 expressed doses of Gnasxl, and 0, 1 or 2 expressed doses of Gnas. We confirm that oedema is due to lack of expression of imprinted Gnas alone. We show that it is the combination of a double dose of Gnasxl, with no dose of imprinted Gnas, that gives rise to the characteristic hyperactive, chunky, oedematous, lethal PatDp(dist2) phenotype, which is also hypoglycaemic. However PatDp(dist2) mice in which the dosage of the Gnasxl and Gnas is balanced (either 2∶2 or 1∶1) are neither dysmorphic nor hyperactive, have normal glucose levels, and are fully viable. But PatDp(dist2) with biallelic expression of both Gnasxl and Gnas show a marked postnatal growth retardation. Our results show that most of the PatDp(dist2) phenotype is due to overexpression of Gnasxl combined with loss of expression of Gnas, and suggest that Gnasxl and Gnas may act antagonistically in a number of tissues and to cause a wide range of phenotypic effects. It can be concluded that monoallelic expression of both Gnasxl and Gnas is a requirement for normal postnatal growth and development.Genomic imprinting results in parent-of-origin-dependent monoallelic gene expression. Early work showed that distal mouse chromosome 2 is imprinted, as maternal and paternal duplications of the region  give rise to different anomalous phenotypes with early postnatal lethalities. Newborns with maternal duplication (MatDp(dist2)) are long, thin and hypoactive whereas those with paternal duplication (PatDp(dist2)) are chunky, oedematous, and hyperactive. Here we focus on PatDp(dist2). Loss of expression of the maternally expressed  Gnas cluster Early work showed that certain chromosomal regions can lead to developmental abnormalities when both copies are exclusively maternally or paternally derived. Distal mouse chromosome 2 was one of the first such imprinting regions described, providing evidence that imprinting must affect expression of some genes according to parental origin Gnas cluster can account for much of the phenotype in both MatDp(dist2) and PatDp(dist2). Gnas is a complex imprinted gene cluster with three promoter regions that give rise to protein coding transcripts Nesp, Gnasxl and Gnas. Each of these transcripts has a unique first exon that splices on to a common set of downstream exons . Nesp is exclusively expressed from the maternal allele and Gnasxl from the paternal allele. Gnas is biallelically expressed in most tissues but is preferentially maternally expressed in some Nesp gives rise to NESP55, neuroendocrine secretory protein 55, Gnas to Gsα, the alpha subunit of the Gs signalling protein and Gnasxl to XLαs, a variant form of Gsα which can also function as the alpha subunit of the heterotrimeric Gs protein. Both XLαs and Gsα can act on adenylyl cyclase to induce cyclic AMP production Gnasxl in offspring result in increased demands on the mother (growth promoting) whereas the effects of maternally expressed Gnas result in fewer demands, in accordance with the conflict hypothesis Misexpression of transcripts at the imprinted am exons . Nesp iGnas cluster is controlled by an imprinting control region (ICR), a germline differentially methylated region called the Nespas DMR that covers the promoter of a noncoding antisense transcript, Nespas Nespas DMR is part of a larger DMR that covers the Gnasxl promoter tm1JopNespas allele , a deletion of the Nespas DMR, results in loss of imprinting of Nesp and Gnas, and down regulation of GnasxlGnas transcript is controlled by a second differentially methylated region, the Exon 1A DMR that lies just upstream of Gnas exon 1 tm1JopGnas allele , a deletion of the Exon 1A DMR, results in loss of imprinted expression of Gnas and so the Exon 1A DMR solely regulates the imprinted expression of GnasNespas DMR must interact with the Exon 1A DMR to control the imprinted expression of Gnas although the mechanism of interaction is not yet clear.Imprinted expression of protein coding transcripts at the Gnasxl but lack expression of Gnas in imprinted tissues and also lack expression of NespGnas in imprinted tissues and two expressed copies of Nesp but lack expression of Gnasxl. PatDp(dist2) mice have greatly diminished expression of Gnas in newborn brown fat consistent with imprinted expression Gnasxl consistent with two expressed doses PatDp(dist2) mice have two expressed copies of Gnasxl results in newborns with severely reduced suckling ability that become inert on the day of birth and most die within within a few days of birth Gnasxl can account for much of the phenotype observed in MatDp(dist2) mice. On the other hand, maternal transmission of a null mutation resulting in loss of Gnas transcript gives rise to neonates with oedema and square shaped bodies, most of which die before weaning Oedsml-matGnas, (Oed), results in gross neonatal oedema and pre-weaning lethality Oed mice) Gnas transcripts in imprinted tissues can account for the oedematous phenotype seen in PatDp(dist2) mice but cannot account for the hyperactivity. Thus hyperactivity may be due to a double dose of Gnasxl alone or to the combined effect of a double dose of Gnasxl and loss of Gnas in imprinted tissues.From studies of targeted deletions and an ENU induced mutation it is evident that both XLαs and Gsα play important roles in growth, behaviour and survival shortly after birth, but NESP55 does not Gnasxl and loss of Gnas gives rise to an early hyperactivity lethal phenotype. Furthermore when the expressed dosage of Gnas and Gnasxl is balanced then PatDp(dist2) are neither lethal nor hyperactive. Thus the dosage of Gnas and Gnasxl needs to be balanced for long term survival and normal activity. Furthermore PatDp(dist2) with balanced dosage but two doses of both Gnasxl and Gnas have normal viability and activity but preweaning growth retardation followed by catch up growth. Thus it may be important to have a single dose of Gnasxl and Gnas for normal development or there may be other genes involved in determining the PatDp(dist2) phenotype.We have used genetic approaches to show the combination of over expression of All mouse studies were carried out in accordance with the guidance issued by the Medical Research Council in “Responsibility in the use of animals in bioscience research (May 2008)”, were approved by the MRC Harwell ethical review committee and carried out under the authority of the UK Home Office Project Licence numbers 30/1518, 30/2065 30/2526, and 30/2642.a, nonagouti locus Gnas cluster.Paternal duplication (PatDp(dist2)) mice were generated by the standard genetic method of intercrossing genetically marked translocation heterozygotes and identifying PatDp(dist2) mice with the aid of markers ΔEx1A (T26H +/+ ΔEx1A) , homozygous for ΔEx1A (T26H ΔEx1A/+ ΔEx1A), heterozygous for ΔNespas (T26H +/+ ΔNespas), and heterozygous for both ΔEx1A and ΔNespas (T26H+ΔEx1A/+ ΔNespas +). PatDp(dist2) that were either heterozygous for ΔEx1A, or homozygous for ΔEx1A, or heterozygous for ΔNespas, or heterozygous for both ΔEx1A and ΔNespas were generated by crossing T26H +/++females with either TH26+/+ ΔEx1A, or T26H ΔEx1A/+ ΔEx1A or T26H +/+ ΔNespas or T26H+ΔEx1A/+ ΔNespas+males. For each cross, reciprocal crosses were also set up to generate PatDp(dist2) with wild-type alleles at ΔEx1A and/or ΔNespas. The genetic background of PatDp(dist2) mice was mixed but with major contributions from C3H/HeH (circa 45%) and the Harwell LL stock (circa 45%). Toe or tail clips were taken from offspring at birth and these served as both biopsies for identification and for genotyping. Offspring were genotyped for paternal duplication, ΔEx1A and ΔNespas as previously described To generate PatDp(dist2) with altered dosing of Gsα and XLαs we developed translocation stocks that were heterozygous for Mice were classified for oedema, chunky appearance, activity, tail kink, and paddle feet, a paddling motion of the front feet, by daily visual observation from birth. Mice were removed from the box and observed for 2–3 minutes.Gnas and Gnasxl and for Actb have been previously described . Transcript levels were measured by phosphoimager analysis and quantified as described previously .Total RNA was extracted and northern analysis carried out as described previously . Riboprobes specific for the unique first exons of Truncal blood from mice up to one week of age was collected using heparinised capillary tubes. Plasma was separated following centrifugation at 750×g for 3 minutes and stored at −80°C. Noradrenaline was measured in duplicate using an ELISA kit (Labor Diagnostika Nord GmbH & Co.KG) according to the manufacturer’s instructions. Blood glucose was measured using an Alphatrak Veterinary Blood Glucose Monitoring Meter Kit (Abbott) according to the manufacturer’s instructions. Insulin and glucagon were measured using an ELISA kit (Mercodia) according to the manufacturer’s instructions.2.Mice were weighed weekly from birth and from 4 weeks the length from the tip of the nose to the base of the tail was measured weekly. Body mass index was calculated using the following equation: weight (g)/(length (cm))http://empress.har.mrc.ac.uk).At nine weeks the mice were individually housed in metabolic cages for 24 h, during which time they had free access to a known amount of food and water. After 24 h, the amount of food and water consumed was measured along with the volume of urine produced. After housing in metabolic cages the mice were returned to their home cage. For a fuller protocol, see EMPRess . Mice were weighed and then individually housed in the equipment for 24 h, with food and free access to water. After testing, the mice were returned to their home cage. Indirect calorimetry enabled measurement of oxygen consumption, carbon dioxide production, respiratory exchange ratio and heat production.At 17 weeks of age the mice were weighed and given a recoverable anaesthetic prior to scanning using a General Electric Medical Systems Lunar PIXImus II X-ray densitometer , which allows the fat and lean mass and bone density to be calculated. Mice were placed in a heated box to aid recovery; once fully recovered, they were returned to their home cage.t test (two-tailed) was used for assessing the results of the growth and metabolism studies and expression levels on northern blots. P values <0.05 were considered significant.Fisher’s exact test was used for comparisons of the incidence of PatDps and the occurrence of phenotypic features. Student’s Gnasxl and severely reduced expression of imprinted GnasGnasxl and Gnas we utilised two mutants, ΔEx1A and ΔNespas. ΔEx1A is a deletion of the Exon 1A DMR, the region that regulates the imprinted expression of Gnas. On paternal inheritance Gnas is completely derepressed on the paternal allele ΔEx1A  show normal survival to birth for they were generated at a frequency of 44.5% (of 231 neonates) which is not significantly different from the expected 50% (P>0.05).The ΔNespas mutant allele is a deletion of the promoter and first exon of Nespas, the ICR for the Gnas cluster ΔNespas Gnasxl is downregulated to about 24% of wild type levels  has two expressed doses of e levels  and GnasGnasxl and Gnas (ΔEx1A were produced to provide PatDp(dist2)2∶1 with two predicted expressed doses of Gnasxl and one expressed dose of Gnas. In Cross 2 PatDp(dist2) mice homozygous for ΔEx1A were generated to produce PatDp(dist2)2∶2 with two predicted expressed doses of Gnasxl and two doses of Gnas. Cross 3 gave rise to PatDp(dist2) mice heterozygous for ΔNespas predicted to have slightly more than one expressed dose of Gnasxl and slightly less than one expressed dose of Gnas, called PatDp(dist2)1∶1. From Cross 4 PatDp(dist2) arose that were compound heterozygotes for ΔEx1A and ΔNespas predicted to have slightly more than one expressed dose of Gnasxl and slightly less than two expressed doses of Gnas, called PatDp(dist2)1∶2. For each cross reciprocal crosses were also set up resulting in PatDp(dist2)2∶0.Four crosses were set up to generate PatDp(dist2) mice with altered dosing of and Gnas , Table 1ΔEx1A were known to express two doses of imprinted Gnas, and PatDp(dist2)2∶1 and PatDp(dist2)1∶1 to express one dose of imprinted GnasGnas were found in newborn brown fat from +/ΔEx1A and PatDp(dist2)2∶2 homozygous for ΔEx1A indicating that PatDp(dist2)2∶2 express two full doses of Gnas (Heterozygotes +/ of Gnas .). The incidence of PatDp(dist2)2∶0 generated using the T26H translocation was not significantly different from that found previously There were no significant differences in the incidence or phenotypes of PatDp(dist2)2∶0 from the reciprocal crosses and so the data from these were pooled for the Control cross . The inGnas cluster which lie 21cM apart (www.informatics.jax.org) . Thus in Cross 1, the cross to generate PatDp(dist2)2∶1, recombinants PatDp(dist2)2∶0 and PatDp(dist2) 2∶2 were found. In Cross 3, the cross to generate PatDp(dist2)1∶1, recombinants PatDp(dist2)2∶0 and PatDp(dist2)0∶2 occurred. In Cross 4, the cross to generate PatDp(dist2)1∶2, recombinants PatDp(dist2)2∶2 arose. All recombinants in Cross 2, the cross to generate PatDp(dist2)2∶2, will be identical and indistinguishable from non recombinant PatDp(dist2)2∶2  to birth.It should be borne in mind that the incidence of PatDp(dist2)2∶1, PatDp(dist2)1∶1 and PatDp(dist2)1∶2 is likely to be an underestimate because of recombination between the TH26 breakpoint on chromosome 2 and the ist2)2∶2 . In the ΔNespas that arose in Cross 3 had a normal phenotype and survived for 6 months. This is probably because in such a mouse the expected levels of Gnasxl expression would be nearly 50% of the wild type level and this appears to be sufficient for viability.Interestingly a recombinant PatDp(dist2)0∶2 homozygous for Gnas and Gnasxl are expected to be in balance and in PatDp(dist2)1∶2 predicted to have one expressed dose of Gnasxl and two expressed doses of Gnas, there was a remarkable increase in survival with most PatDp(dist2)2∶2, PatDp(dist2)1∶1 and PatDp(dist2)1∶2 reaching weaning (Gnasxl is disadvantageous but overexpression of imprinted Gnas is not.In agreement with earlier work PatDp(dist2)2∶0 generated using the T26H translocation only survived for a few days  weaning . After wOed mutant, a point mutation in Gnas exon 6 At birth the PatDp(dist2)2∶0 mice had the typical phenotype . Thus, mGnas, thus PatDp(dist2)2∶1, PatDp(dist2)2∶2, PatDp(dist2)1∶1 or PatDp(dist2)1∶2 were oedematous 2∶1, a number were noted to be chunky , indicatGnasxl together with loss of expression of imprinted Gnas gives rise to this phenotype. However, if the double dose of Gnasxl is matched with a double dose of Gnas the chunky phenotype was not seen. For normality the expressed doses of Gnasxl and Gnas need to be balanced 2∶1, PatDp(dist2)2∶2, PatDp(dist2)1∶1 and PatDp(dist2)1∶2 described as chunky compared to PatDp(dist2)2∶0. In addition, fewer PatDp(dist2)2∶2 and PatDp(dist2)1∶1 were chunky compared to PatDp(dist2)2∶1 . Overallbalanced .Gnas and Gnasxl have the same predicted expressed dosage in PatDp(dist2)1∶1 as in wild type. Loss of functional imprinted Gnas in the presence of a single expressed dose of Gnasxl is not associated with hyperactivity in the first few days of life Gnas knockout model Gnasxl and loss of Gnas in imprinted tissues Gnasxl combined with loss of imprinted Gnas expression is responsible for the hyperactivity phenotype.Hyperactivity in PatDp(dist2)2∶0 although manifested on the day of birth, becomes fully developed by the next day . There wGnasxl together with either loss of, or a single dose of imprinted Gnas gives rise to this phenotype. For normality the expressed doses of Gnasxl and Gnas need to be balanced 2∶0, are not significantly different in PatDp(dist2)2∶1 but are reduced in PatDp(dist2)2∶2, PatDp(dist2)1∶1 and PatDp(dist2)1∶2. Furthermore, fewer PatDp(dist2)2∶2 and PatDp(dist2)1∶1 exhibited either tail kink and/or paddle feet than PatDp(dist2)2∶1 . Overallbalanced .Gnas in tissues in which it is imprinted, chunkiness, hyperactivity, tail kink and paddle feet to overexpression of Gnasxl, combined with loss of imprinted Gnas expression.Thus oedema can be attributed to lack of expression of ΔEx1A whereas nonduplication littermates of PatDp(dist2)2∶2 are heterozygous +/ΔEx1A and thus have biallelic Gnas expression. We tested neonates for glucose and key hormones involved in energy homeostasis .Studies of metabolism were undertaken in PatDp(dist2)2∶0 and and PatDp(dist2)2∶2 and their nonduplication litttermates. The nonduplication littermates of PatDp(dist2)2∶0 are wild type for eostasis . In PatDΔEx1A the levels of glucose, insulin and glucagon and noradrenaline were not significantly different from wild type indicating that overexpression of Gnas is not associated with altered glucose metabolism.In newborn +/ΔEx1A sibs were carried out. In addition studies of +/ΔEx1A and their +/+ sibs from a cross of C3H/HeH females to +/ΔEx1A males were undertaken. For weight studies between birth and weaning the weights of both sexes were combined as no significant differences in the weights of males and females were found, and after weaning only males were used. Mice were weighed on the day of birth and weekly thereafter.Further studies from birth to adulthood of PatDp(dist2)2∶2 and their nonduplication +/ΔEx1A mice were lighter at birth, 90.1% (P = 0.021), of the weight of their wild type siblings, indicating prenatal growth retardation. Growth retardation became more pronounced during the first week of life but by weaning +/ΔEx1A had started to catch up so that by nine weeks their weight was no longer significantly different from wild type . Similar findings of prenatal and preweaning growth retardation followed by catch up growth has been reported on paternal inheritance of the Ex1A-T-CON allele which also results in loss of imprinted expression of GnasThe weights are shown as a percentage of their wild type siblings . The +/Δld type . SimilaΔEx1A/++ΔEx1A so all the nonduplication sibs were heterozygous for ΔEx1A. PatDp(dist2)2∶2 did not differ in weight from their nonduplication +/ΔEx1A sibs at birth or at one week of age indicating that PatDp(dist2)2∶2 were also growth retarded and to the same extent as +/ΔEx1A. By two weeks of age PatDp(dist2)2∶2 were even lighter in weight than +/ΔEx1A and continued to weigh less than +/ΔEx1A until 6 weeks but subsequently did not differ significantly in weight from +/ΔEx1A. Thus postnatal growth retardation in PatDp(dist2)2∶2 peaked at three weeks and was followed by catch up growth (PatDp(dist2)2∶2 were generated by Cross 2, T26H++/+++ X T26H+p growth .Gnas in +/ΔEx1A results in growth retardation before weaning resulting in proportionately smaller mice followed by catch up growth in adulthood. PatDp(dist2)2∶2 with predicted balanced expressed dosage of Gnas and Gnasxl show a more severe pre-adult growth retardation resulting in proportionately smaller mice followed by catch up growth by 10–11 weeks of age and overall increased weight thereafter.Thus loss of imprinting of ΔEx1A mice, their wild type sibs, and PatDp(dist2)2∶2 and their +/ΔEx1A sibs was measured weekly from 4 weeks. +/ΔEx1A mice were shorter (93.2±0.5% s.e.m of the length of wild type sibs) than their wild type sibs at 4 weeks  but caught up in length by 10 weeks. PatDp(dist2)2∶2 were even shorter than +/ΔEx1A from 4 to 6 weeks  but thereafter did not differ in length from +/ΔEx1A.The body length of +/ΔEx1A than wild type at 4 weeks  but did not differ from wild type thereafter. No difference in BMI was found between +/ΔEx1A and PatDp(dist2)2∶2 (n = 10–12).Body mass index, BMI was significantly lower in +/ΔEx1A and wild type sibs (n = 4–14) or in +/ΔEx1A and PatDp(dist2)2∶2 (n = 10–12) (data not shown).Analysis of 24h food intake at 10 weeks showed no difference in +/ΔEx1A and wild type sibs (n = 5–12) or in +/ΔEx1A and PatDp(dist2)2∶2 (n = 9) (data not shown).The results of DEXA analysis at 17 weeks indicated there were no significant differences in fat or lean mass in +/ΔEx1A tended to show increased oxygen consumption and carbon dioxide output in comparison with wild-type littermates and these increases were statistically significant (P<0.03) by 21 weeks of age in +/ΔEx1A mice indicative of a raised metabolic rate (At 10 weeks of age +/lic rate .Gnas as oedema was not seen in PatDp(dist2) in which Gnas expression was restored to normal. This agrees with the results from other studies indicating that null or loss of function mutations of Gnas are associated with neonatal oedema Gnas expression was restored to normal Oed even more so with oedema declining somewhat in both genetic conditions before birth in utero could conceivably affect survival to birth and so the restoration of imprinted Gnas expression in PatDp(dist2)2∶1 and PatDp(dist2)1∶1 may account for their increased incidence at birth compared to PatDp(dist2)2∶0.The oedema in PatDp(dist2)2∶0 is likely to be due to loss of expression of imprinted Gnasxl combined with loss of imprinted Gnas expression. A considerable improvement in the incidence of newborns that were neither hyperactive nor chunky and some improvement in survival to seven days was found when biallelic expression of Gnasxl was combined with normal monoallelic expression of Gnas in PatDp(dist2)2∶1. However, for full viability and loss of hyperactivity, biallelic expression of Gnasxl needs to be balanced with biallelic expression of imprinted Gnas or monoallelic expression of Gnasxl with monoallelic expression of Gnas. This need for balanced dosage is in accord with observations indicating that maternally expressed Gnas and paternally expressed Gnasxl act antagonistically The other neonatal phenotypes of PatDp(dist2), namely lethality, hyperactivity, paddle feet, chunky appearance, tail kink, can be attributed to biallelic expression of Gnasxl combined with biallelic, and thus overexpression of imprinted Gnas, in either PatDp(dist2)1∶2 or +/ΔEx1A results in normal viability. Normal viability was also recently reported for the Ex1A-T-CON mutant in which Gnas is overexpressed Gnas has less drastic results on phenotype than overexpression of Gnasxl. But in the presence of two expressed copies of Gnasxl it is critical to have two expressed copies of Gnas in order to survive to weaning.Interestingly monoallelic expression of mΔNesp55 neonates Gnasxl and no expressed doses of imprinted Gnas. The hyperactivity in PatDp(dist2)2∶0 and mΔNesp55 may reflect a central neurological deficit. The finding of raised noradrenaline in PatDp(dist2)2∶0 together with a recent report showing that Gnasxl is expressed in neonatal muscle Gnasxl KO pups might be at least partly due to the lack of XLαs in muscle.PatDp(dist2)2∶0 are hyperactive and hypoglycaemic and both these phenotypes may contribute to their early death. Hyperactivity has also been reported for some mΔNesp55 neonates are also hypoglycaemic and although poor feeding may well contribute to their hypoglycaemia they may, in addition, have a defect in glucose counterregulation Gnasxl and Gnas, glucose metabolism is restored to normal, suggesting that both Gnasxl and imprinted Gnas have a role in neonatal glucose metabolism.PatDp(dist2)2∶0 are hypoglycaemic with low insulin and raised noradrenaline. Gnasxl proteins to other aspects of the PatDp(dist2)2∶0 phenotype is as yet unknown.Both Gsα and XLαs have several variants. Thus C-terminally truncated neural specific forms GsαN1 and XLN1 of Gsα and XLαs respectively are known Gnas and Gnasxl have roles in the phenotype of PatDp(dist2). Loss of the orthologous GNAS transcript in humans is associated with a number of endocrine and bone disorders including pseudohypoparathyroidism, Albright Hereditary Ostedystrophy and progressive osseous heteroplasia GNAS and pseudohypoparathyroidism type Ib is caused by defective imprinting at the cluster  mice hyperactivity commencing shortly after birth and elevated noradrenaline are also features of raised Gnasxl levels. It would be of interest to ascertain if metabolism and activity in PHP-Ib patients are affected in the early postnatal period and if so, whether elevated GNASXL is responsible.Our data indicate that both Gnas in +/ΔEx1A results in postnatal growth retardation followed by catch up growth and can be attributed solely to loss of imprinting, resulting in biallelic expression, of Gnas. The effects on postnatal growth in +/ΔEx1A parallel the growth pattern seen in +/Ex1A-T-CON mice which also have loss of imprinting of GnasGnas clearly has a role in growth. Paternally expressed Gnasxl is also known to affect postnatal growth and mutations resulting in loss of functional Gnasxl also result in postnatal growth retardation Gnas and underexpression of Gnasxl results in growth retardation accords with the parental conflict theory that predicts that maternally expressed imprinted genes will be growth inhibiting and paternally expressed imprinted genes will be growth enhancing Gnas and Gnasxl. However PatDp(dist2)2∶2 mice show a more extreme postnatal growth retardation than seen with overexpression of Gnas in +/ΔEx1A. Thus biallelic expression of both Gnas and Gnasxl has resulted in postnatal growth retardation. One interpretation is that it is important to have monoallelic expression of antagonistic Gnasxl and Gnas for normal growth. The advantages of monoallelic expression of both Gnasxl and Gnas must outweigh the costs associated with functional haploidy.We have shown that loss of imprinting of Figure S1Expression of Gnasxl was reduced on paternal inheritance of ΔNespas. (A) Northern blot of Gnasxl and β-actin loading control using 2.5 µg of poly (A)+ RNA from 15.5-dpc embryos. MatDp(dist2) have no expressed copies of Gnasxl, whilst PatDp(dist2) have two expressed copies, leading to the absence of the 2.5 kb band and the presence of a strong 2.5 kb band respectively. (B) Bar chart showing the Gnasxl expression levels in +/ΔNespas (mean ± s.e.m 24±3.56%) were decreased on comparison to wild-type (+/+) (mean ± s.e.m 103±9.39%) *P = 0.005 . The mean ± s.e.m was calculated for 3 wild-type (+/+) and four +/ΔNespas.(TIF)Click here for additional data file.Video S1Hyperactivity in PatDp(dist2). Video of a 6 day old PatDp(dist2) and wild-type littermate showing almost continual activity and rapid movement with paddling of the front feet in the PatDp(dist2) but very little movement in the wild-type. The PatDp(dist2) has the characteristic tail kink or bend, and by 6 days is clearly smaller than the wild-type but oedema and a chunky appearance are no longer evident. The mice have a genetic background that is 50% Mus musculus castaneus and 50% laboratory mouse. On this background there was slightly better survival of PatDp(dist2) to 7 days (10/23) compared with 3/28 on an entirely laboratory mouse background  but no difference in survival to weaning.(MP4)Click here for additional data file.Video S2Hyperactivity in PatDp(dist2). Video of the same PatDp(dist2) and wild-type littermate at 6 days as in Video S2 and the PatDp(dist2) at 5 days of age on its own. The video shows the ability of the PatDp(dist2) to right itself after falling on its back, as well as its continual activity and rapid movement.(MP4)Click here for additional data file.