Activating PAX gene family paralogs to complement PAX5 leukemia driver mutations

Analysis of the 5’ enhancer/promoter regions of both PAX2 and PAX5, along with their intronic and exonic DNA, indicated numerous iterations of the consensus (TGGAAANNYNY) TonE binding site (S9A and S9B Fig) [47]. To determine whether NFAT5 was involved in hyperosmolarity-induced expression of PAX2 and PAX5 and to concurrently assess whether such PAX upregulation directly affected downstream gene modulation, we performed siRNA knockdown of these three genes (Fig 8B and 8C). We found that siRNA knockdown of NFAT5 was sufficient to abrogate PAX2 upregulation in response to 80mM K-gluconate in Reh cells (Fig 8C). Similarly, knockdown of NFAT5 quenched hyperosmolarity mediated increases in the solute carriers, SLC5A3 and SLC6A6, both of which are known targets of NFAT5 (S8B Fig) [48]. Interestingly, neither PAX5 nor PAX5 downstream genes upregulated in response to hyperosmolarity were affected by NFAT5 knockdown (Fig 8C), consistent with a separate, NFAT5 independent mechanism for induction of PAX5 or, at least, reduced sensitivity of PAX5 to changes in NFAT5 levels. Importantly, knockdown of PAX5 itself led to reductions in expression of the downstream genes we assessed, while siRNA directed against PAX2 had little effect (Fig 8C), suggesting that hypertonic induction of residual wild type PAX5 expression outweighs PAX2 with respect to regulation of their common targets. We note that PAX2 expression is detectable as a transcript, but insufficient to measure at the protein level by western blot.

The PAX5 mutation in Reh cells creates a frameshift leading to premature termination and is thus expected to be subject to nonsense-mediated decay. However, western blot indicates that, in addition to a full-length PAX5 protein corresponding to the wild type allele, a truncated polypeptide that is likely non-functional is apparently generated from the mutant allele, albeit at reduced abundance, suggesting that nonsense-mediated decay is incomplete (as evident in Fig 8B, where both products are specifically targeted by siRNA directed against PAX5). Reh cells should therefore contain mRNA from both the wild type and mutant PAX5 alleles. To determine if either salt treatment differentially activates the wild type as opposed to the mutant PAX5 allele in Reh cells, we analyzed RNA-seq data and compared the total number of reads obtained from each allele (and that also include the distinguishing mutation). In untreated cells, 27 of 105 total, non-normalized reads (26%) corresponded to transcripts from the mutant allele. In K-gluconate or CaCl2 treated cells, the equivalent proportions of mutant transcripts were 54/261 (21%) and 36/135 (27%), respectively. (Using a two-tailed test to compare two population proportions, for untreated versus K-gluconate treated cells, the Z-score is 1.05 and p-value is 0.29. The same comparison for CaCl2 treated cells yields a Z-score of -0.17 and p-value of 0.87). These differences are not significant. We conclude that neither salt treatment discriminates between wild type and mutant allele when activating PAX5 expression, as reflected in proportionately increased total read counts.

As numerous studies have shown, long term, in vitro, cell culture inherently selects for gene expression profiles differing from those seen for primary tissue samples [49,50]. To further evaluate whether the PAX2/5 response to hyperosmolarity is one that is intrinsic to ALL cells both in vitro and in vivo, we screened 10 primary pre-B ALL samples for PAX5 mutations, using Sanger DNA sequencing. Of those samples, one, from a 19 year-old male with trisomy 21 Down syndrome, possessed a heterozygous p.(K198Qfs*44) mutation, resulting in frameshift leading to early stop and protein truncation (see Methods). Pre-B ALL occurs more commonly in Down syndrome individuals and is felt to be biologically distinct from disease occurring in non-Down syndrome patients [51]; nevertheless, inactivating mutations of PAX5 are detected at similar frequency in Down syndrome-associated pre-B ALL [52]. Due to limited sample availability from this patient, we performed a single test employing primary cells alongside multiple replicates using primary cells expanded through passage in mice (see Methods). Whether direct from the patient or passaged through mice, 24 hour exposure to 80mM K-gluconate resulted in increased expression of PAX5, as well as several but not all downstream targets seen previously with Reh and 697 cells (Fig 9A and S10A Fig). PAX2 expression was not detected in this assay; however, this may be due in part to low RNA input levels, which were constrained by sample quantity.

The osmotic concentrations of K-gluconate or CaCl2 we evaluated in vitro would prove lethal if administered clinically. However, mannitol is also known to activate NFAT5 [53] and is used to adjust serum hyperosmolar concentrations to high levels in certain clinical settings [54]. To test whether mannitol could be employed to upregulate PAX2 or PAX5 in pre-B ALL, we treated Reh cells with 80mM or 160mM mannitol for 24 hours, prior to FACS sorting for live cells and harvesting of RNA. qRT-PCR demonstrated dose-dependent increases both for PAX2 and especially for PAX5, along with similar changes in downstream gene expression, albeit not to the level seen with K-gluconate (Fig 9B and 9C). Importantly, 160mM is near the range of clinically achievable therapeutic concentrations for mannitol [54]. Comparison of 160mM mannitol with 80mM K-gluconate or CaCl2, followed by FSC-A/SSC-A sorting of live cells and subsequent measurement of culture expansion demonstrated slightly reduced growth potential for K-gluconate and CaCl2 treated cells as compared to cells grown in 160mM mannitol or normal media (S10B Fig). Interpretation of long term viability in response to hyperosmolarity was complicated due to the noticeably brief induction of PAX2/5 (Fig 5E), coupled with the generally harsh nature of such treatment, even with only 24 hour exposure. The growth delay observed with K-gluconate in this case may largely be due to cell cycle arrest or other adverse effects of elevated hyperosmolarity [55], rather than the PAX dependent, developmentally programed exit from the cell cycle we appeared to observe with continuous PAX re-expression. However, even in vitro, mannitol appears to be better tolerated, and thus it or related organic osmolytes may present options for modulating tonicity that could prove tolerable in vivo.

Liu et al. recently demonstrated that restoration of PAX5 expression can reverse the developmental blockade holding PAX5-mutated pre-B ALL cells in a continuously replicating, developmentally immature state [18]. We have confirmed that result and extended it further by showing that PAX5’s closely related paralogs, PAX2 or PAX8, neither of which is mutated in ALL nor ordinarily expressed in lymphocytes, can function equivalently to normalize differentiation and growth of pre-B ALL cells. Moreover, we have shown that endogenous PAX2 expression, and unexpectedly also PAX5 itself, can be upregulated to promote similar effects on differentiation of pre-B ALL cells under hypertonic conditions.

While germline loss-of-function mutations are a cause of familial pre-B ALL [13,14], demonstrating that PAX5 deficiency can ultimately initiate leukemogenesis, loss of PAX5 activity is not by itself sufficient, and development of leukemia requires additional cooperating mutations. Cancer genome sequencing has identified a wide diversity of mutations [8,9], such that no two ALL patients are likely to share identical mutational profiles. Reh and 697 cells, tested here as well as by Liu et al. [18], are quite dissimilar, with 697 cells having only a few coding sequence alterations while Reh cells have considerably more, with very little overlap (S11 Fig). In particular, Reh cells contain a heterozygous loss-of-function PAX5 mutation [33], whereas in 697 cells, PAX5 is intact (per our sequencing, S13 Fig). However, an upstream regulator of PAX5, E2A (TCF3), is at least partially inactivated via a translocation involving PBX1 [35], suggesting that there may be similarly reduced expression of PAX5 in 697 cells. Regardless, targeting PAX2/5/8 activity may prove beneficial even in those patients lacking PAX5 mutations. Liu et al. also demonstrated that PAX5 replenishment succeeded in curing a transgenic mouse model of ALL, driven by PAX5 knockdown combined with Stat5 activation [18]. The fact that PAX5 re-expression normalizes growth and differentiation in pre-B ALL with divergent genetic backgrounds and mutational signatures, including with Down syndrome associated ALL as tested in primary cells, suggests that even after cooperating mutations have arisen, loss of PAX5 activity continues to support the leukemic state.

Consistent with the concept of oncogene addiction, in which secondary mutations are dependent upon driver mutations for maintaining the cancer phenotype [56], acquisition of additional mutations may therefore possibly render ALL cells even more vulnerable following replacement of PAX5 activity.

Current approaches for treatment of pre-B ALL continue to rely on chemotherapy and, more recently, immunotherapy. Chemotherapy is often successful in pediatric settings [1] but is associated with considerable toxicity, long-term side effects [2], and substantially reduced efficacy in older children and adults, where allogenic stem cell transplant is more heavily relied upon [3]. Recent breakthroughs in CAR T cell therapy have shown great promise in treating certain disease presentations, specifically those which are highly CD19 positive [57]. However, hurdles remain, including clonal selection for PAX5 deletion with consequent downregulation of the CD19 target antigen, leading to disease resistance [58]. Since CD19 is a direct target of PAX5 and, as we have shown, can be activated equally well by PAX2 or PAX8, the therapeutic approach contemplated here may work in conjunction with CAR T cell therapy by increasing levels of the targeted CD19 B cell antigen, even after loss of PAX5. CD19 can also be targeted through other forms of therapy, such as with antibody-drug conjugates [59].

Our observations demonstrate the use of gene paralogs to resolve a human disease phenotype. A remaining challenge, however, involves approaches for activating developmentally silenced genes in vivo. We tested a variety of compounds based upon previously described properties as either generally reversing repressive chromatin modifications (zebularine, hydralazine, valproic acid, azacitidine, and vorinostat) or as mechanistically undefined inducers of lymphocyte differentiation (ATRA, methotrexate, and phorbol 12-myristate 13-acetate (PMA)). None consistently activated PAX2 or PAX8 expression or otherwise promoted pre-B ALL cell differentiation under conditions we evaluated.

Another class of compounds we tested affect cell membrane potential, the modulation of which has been shown in model systems to induce a variety of developmental transcription factors, including, for example, PAX6 [40]. We observed induction of PAX2, but not PAX8, as well as increases in downstream differentiation markers in response to K-gluconate. After testing a variety of other salts as well as several non-ionic modulators of cell membrane potential, we concluded that our observation was likely a cellular response to hypertonicity. During water diuresis, physiological concentrations of salts, mainly NaCl, in the renal inter-medullary interstitial fluid reach concentrations ranging from 600 to more than 1000mOsmol/kg H2O. Interestingly, survival mechanisms for cells in these conditions include the anti-apoptotic upregulation of PAX2, which has been shown to peak in mouse intermedullary collecting duct cells at ~500mOsmol/kg H2O [30], similar to what we observed in Reh cells (~400-540mOsmol/kg H2O in RPMI media). Unexpectedly, we observed that hypertonicity also induced expression of PAX5 in pre-B ALL.

RNA-seq performed in conjunction with GSEA highlighted similarities and differences resulting from expression of PAX2 or PAX8, compared to PAX5, in PAX5-deficient ALL cells. In a pairwise comparison of any of the three PAX factors, slightly fewer than half of all gene sets exhibiting significant expression changes were common to both, and only 13% of all gene sets (57/440, Fig 6A) enriched by PAX5 were commonly modulated by all three PAX genes. Importantly, however, the group of gene sets commonly regulated by all three PAX factors includes PAX5 targets most relevant to B cell maturation (Fig 6D), consistent with our findings that all three PAX factors similarly promote differentiation of PAX5-deficient ALL cells. We speculate that PAX5 target genes likely reside in accessible chromatin configurations in pre-B cells, such that even imperfect PAX activity from a paralog may readily induce their expression. In contrast, gene sets exhibiting significant enrichment following treatment with CaCl2 or K-gluconate exhibited much greater overlap with PAX5, and a majority of gene sets showing enrichment with PAX5 (221/420, Fig 6B) or that were commonly enriched by all three PAX factors (43/57, Fig 6C) were also enriched after treatment with CaCl2 or K-gluconate. This may not be surprising given that treatment with either salt induced expression of PAX2 and, especially, PAX5 itself. Finally, it is worth emphasizing from a translationally relevant standpoint, that a set of 31 genes found by Liu et al. to undergo significant regulation during ALL remission, as induced by Pax5 restoration in a mouse model of Pax5-deficient ALL, were similarly modulated by all tested conditions in our studies, whether it be PAX5, PAX2, PAX8, K-gluconate, or CaCl2 (Fig 6C).

We found that components of the NFAT5 pathway, including NFAT5 and TNFS13B (BAFF), along with its receptor, TNFRSF13C (BAFF-R), are upregulated in response to many or all of our treatments (i.e., PAX2/5/8 or salt treatment, Fig 8A and S1 Dataset). Named “nuclear factor of activated T cells 5,” for its role as a transcriptional coordinator of T cell immune response [60], NFAT5 is the only known osmosensing mammalian transcription factor and is active in a variety of cell types, including B cells [45,46]. Indeed, siRNA mediated knockdown of NFAT5 in Reh cells led to a reduction in PAX2 expression in response to hyperosmolarity (Fig 8B and 8C). However, the added observation that PAX5 expression was not affected by NFAT5 knockdown suggests either the presence of a separate, non NFAT5 related, osmosensing pathway upstream of PAX5, or alternatively, a substantially lower threshold for NFAT5 abundance to achieve upregulation of PAX5 under these conditions. In support of the latter, PAX5 appears to contain more potential NFAT5 binding sites than PAX2 (S9 Fig). Separately, these siRNA experiments showed that PAX5 upregulation had a greater effect on downstream gene regulation, and presumably B cell maturation, than did PAX2 (Fig 8B and 8C). Based on our observations from earlier experiments (Figs 2–4), where PAX2 effectively functionally mimicked PAX5, and the substantially lower level of PAX2 expression present relative to the induced levels of PAX5 in response to hyperosmolarity (~20 fold), we believe this most likely reflects relative levels of expression, rather than differences in functionality.

Given that components of the hypertonicity response pathway are highly conserved from single cell organisms to mammals [61], it seems reasonable to speculate that PAX genes, including PAX2 and PAX5, may play a role in osmotic adaptation across various tissue types. In fact, similar to our observations, upregulation of PAX2 occurs in mouse embryonic fibroblasts in response to hypertonicity [48]. Secondary lymphoid organs, including spleen and thymus, maintain a remarkably high osmolar environment compared to serum and other tissue [62]. It should not be overlooked that a decrease in cell size, which we observed upon expression of PAX2/5/8, normally accompanies the large-to-small pre-B cell transition as cells begin their migration from the bone marrow to secondary lymphoid organs. It is possible that exposure to differences in local osmolarity across these compartments could play a role in normal lymphocyte development. Whether upregulation of PAX2 and/or PAX5 is a normal physiologic response to osmotic stress in lymphocytes or a vestigial pathway more heavily relied on in other tissues such as the kidney, but which is capable of artefactual activation under extreme circumstances, we show here that osmotic stress exposes a potential therapeutic target for activating elements of the normal B cell differentiation program.

The osmolar concentration required for peak induction of PAX2 and PAX5 is, just barely, outside the clinically achievable range for serum based on maximum recommended dosing for mannitol [54]. It is possible that specialized delivery methods, manipulation of dosage levels, and/or exposure time may bridge this gap. It is also worth noting that serum osmolar concentrations within this range are sometimes encountered in acutely ill diabetes mellitus patients with hyperglycemic hyperosmolar syndrome [63]. However, even if the highly hyperosmolar conditions we subjected ALL cells to in vitro are not therapeutically tenable in vivo, they do suggest that the complexity of kinases and other components of the signaling pathway responding to hypertonicity, including those regulating NFAT5, at least in the case of PAX2 [64], may be ripe for investigation as drug targets.

An additional limitation relates to duration of therapy, as the replacement of PAX5 activity may only have a temporary effect on differentiation of ALL cells, though this may still be beneficial either as a form of induction therapy or as an adjuvant when combined with CAR T cell, other therapies targeting CD19, or conventional chemotherapy. Intriguingly, a relevant recent in vitro study demonstrated that hyperosmotic stress achieved with salt or mannitol treatment synergized with chemotherapeutic drugs to kill ALL cells via an NFAT5 dependent mechanism, although activation of PAX genes was not investigated [53]. It should also be emphasized that remissions achieved with differentiation therapy employing ATRA for promyelocytic leukemia can actually be enduring [7]. Finally, if differentiation of pre-B ALL cells could be pushed as far as to the plasma cell stage, where PAX5 expression is normally extinguished [65], then mutations inactivating PAX5 could become inconsequential, anyway.

Finding the right balance of PAX gene expression is another issue. PAX2, when activated, can behave as an oncogene in solid tumors [66], and PAX5 is normally down-regulated during plasma cell differentiation [65]. However, our RNA-seq data suggest that there may be an auto-regulatory ceiling for PAX gene expression, particularly for PAX5. Specifically, by examining total PAX5 transcripts and comparing differences in the read ratios of SNPs discriminating between native and exogenous PAX5, we observed an apparent suppression of endogenous PAX5 transcript by PAX5 transgene expression, and to a lesser extent, by the expression of PAX2 or PAX8 transgenes (S12 Fig). Of course, unless PAX gene activation is confined only to the leukemic population of cells, there may be undesirable effects in other tissues, although compared to oncogenic mutations, PAX gene activation by osmoresponsive mechanisms is unlikely to be permanent. Moreover, some current cancer therapies employ treatment with epigenetic modifier drugs, such as azacitidine, capable of producing genome-wide and persistent activation of many genes across multiple tissues [67].

The strategy implemented here, to activate expression of intact and functionally similar paralogs of mutated cancer-driver genes to therapeutically restore cellular differentiation, could potentially be extended to other types of cancer. For example, inactivating RUNX1 mutations frequently occur in acute myeloid leukemia, where upregulation of RUNX2 or RUNX3 exhibits anti-leukemic effects [68]. More generally, a wide variety of non-cancer illnesses possess etiologies for which complementation of inactivating mutations by activating gene paralogs may prove useful, extending the potential therapeutic application of this concept. For example, in spinal muscular atrophy, causative loss-of-function mutations in SMN1 can be rescued by a recently approved therapy which uses an antisense compound to promote exon retention in an alternatively spliced yet otherwise identical paralog, SMN2 [69]. Finally, hypertonic activation of PAX gene expression offers an example of emerging “electroceutical” approaches based on manipulation of biophysical phenomena [41].

Leukemia cells were collected, after informed consent, through the Cell Bank of the Center for Cancer and Blood Disorders at Children’s Hospital Colorado. The Cell Bank protocol is approved by the Colorado Multiple Institutional Review Board (COMIRB #00–206).

Animal use was approved by the Animal Care and Use Committee of the University of Colorado Denver (Protocol 66912(12)1E).

Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Marshall Horwitz ([email protected]).

Reh cells (female [32]) and HEK293T cells (female [70]) were obtained from ATCC. 697 cells (male [34]) were obtained from DSMZ. As suggested by their supplier, Reh and 697 cells were grown in 10% FBS/RPMI 1640 (Gibco, 11875–093), and HEK293T cells were grown in 10% FBS/DMEM (Gibco, 11995–065). All cells were grown at 37°C, under 5% CO2 and were passaged every 3–4 days at confluencies suggested by their suppliers. Cell aliquots were cultured for no more than ~20 passages. Based on hybrid capture exome sequencing, 697 cells are reported in the Cancer Cell Line Encyclopedia (CCLE) [71] as being heterozygous for the PAX5 null mutation p.R225fs; however, we could not confirm this mutation upon Sanger DNA sequencing (S13 Fig), and review of the CCLE primary data shows that both read depth and variant allele count at this position are minimal, consistent with artifact. To verify the identity of the 697 cell line in our possession, we performed Sanger DNA sequencing and confirmed the existence of two separate and uncommon CCLE-reported gene mutations, intrinsic to this cell line; heterozygous NRAS p.G12D and hemizygous GPR112 p.D2657del (S13 Fig). While not identified in the CCLE, Reh cells have been previously reported to contain a heterozygous p.A322fs mutation [33], which we confirmed by Sanger sequencing (S13 Fig). For further verification of cell line identity, low passage samples of both Reh and 697 cells were submitted to ATCC for short tandem repeat profiling and validation by comparison with known profiles for Reh and 697 cell lines. The samples were profiled using the PowerPlex 18D system (Promega), which provides a signature encompassing 17 short tandem repeat loci plus Amelogenin (S14 and S15 Figs).

Ten B ALL samples were screened for mutation in PAX5 by Sanger DNA sequencing. One sample, CHCO-7657, was found to contain a heterozygous PAX5 mutation resulting in p.K198fs (S16 Fig). A subset of primary cells were stored in liquid nitrogen and additional leukemia cells were passaged through NOD scid gamma (NSG) mice once and amplified in NOD scid gamma Il3-GM-SF (NSGS) mice (both purchased from Jackson Labs), prior to storage in liquid nitrogen. Recipient mice were irradiated with 200cGy via X-irradiator prior to leukemia injection. Prior to experimentation, cells were thawed and resuspended in 20%FBS/MEM-alpha (Gibco,12561–056) which had been preconditioned with OP9 feeder cells (ATCC) seeded the day prior at 3×105 cells/T75 flask, in 10mL media. Primary cells were overlaid onto and co-cultured with these OP9 feeder cells for 24 hours, followed by treatment with 80mM K-gluconate for an additional 24 hours before sorting for live cells and isolation of RNA.

Lentiviral plasmids pRRL-CMV-IRES-hrGFPII and pLVX-EF1α-IRES-ZsGreen, were obtained from the University of Washington Diabetes Research Core and Clontech, respectively. PAX5 isoform 1 cDNA [13] was utilized for these studies. PAX2b (catalog # SC300041) and PAX8a cDNA (SC122658) were obtained from OriGene. The PAX2b isoform was chosen due to its relatively abundant expression in human tissues and close resemblance to full length PAX5 (S1 Fig). PAX8a is the most commonly expressed isoform [72] but includes an extra serine-rich ~60 amino acid region between the partial homeodomain and transactivation domain that is absent from PAX2 or PAX5. EcoR1 sites were added to both ends of cDNA upon amplification with primers (PAX2F: 5’ACAGTAGAATTCGCCACCATGGATTACAAGGATGACGACGATAAGATGGATATGCACTGCAAAGCAGACC, PAX2R: 5’CTAGTGGCGGTCATAGGCAG, PAX8F: 5’ACAGTAGAATTCGCCACCATGGATTACAAGGATGACGACGATAAGATGCCTCACAACTCCATCAG, PAX8R: 5’CTACAGATGGTCAAAGGCCG) and were employed for ligation into plasmids. PAX5p.V26fs was created via primer directed mutagenesis (Forward primer: 5’[Phos]GGGGTTTTTGTGAATGGACGG, Reverse primer: 5’[Phos]CCCCAAGCTGATTCACTCCTCC). Directionality and integrity were confirmed by Sanger sequencing. Expected protein size/expression was verified by western blot in HEK293T cells (visible in S3 Fig). Lentivirus production was performed by Allele Biotech and the University of Washington Diabetes Research Core Facility.

Cells were transduced using “spinoculation.” Briefly, cells were washed twice in serum-free RPMI and distributed at 2×105 cells/well in a 96-well plate. Lentivirus (MOI 10–20) was added, along with 4.5 μg/mL polybrene for a total of ~100 μL volume/well. Cells were centrifuged at 1200×g at 30°C, for 2 hours. Supernatant was then removed and cells resuspended in a total of 1 mL growth media, then analyzed at day 4 post-transduction unless otherwise noted.

Cells were lysed using RIPA buffer with complete protease inhibitor (Roche), 1mM Na3VO4 and 1mM PMSF. Lysates were quantified with the Pierce BCA Protein Assay Kit (ThermoFisher) and electrophoresed on MINI-protein TGX gels (BioRad) and transferred onto PVDF membrane (BioRad). Membranes were immunoblotted with indicated antibodies. All primary antibodies were diluted in 1% milk or BSA in TBST and were incubated overnight at 4°C. Blots were incubated in secondary antibody for 1 hour at room temperature, also in 1% milk or BSA.

Flow cytometry for cell surface markers was performed on a BD LSR II flow cytometer using indicated antibodies. For antibody staining, cells were washed twice in sorting buffer (1%FBS/PBS), prior to incubation in antibody (diluted in sorting buffer) on ice and in the dark, for 30 minutes. Cells were then again washed twice in sorting buffer, and resuspended in 300–500μL sorting buffer prior to analysis. All staining and washing was done in 96 well, flat bottom plates. In between washes, cells were spun down for 3’ at 300×g. Plates were overturned and shaken to remove buffer. FACS was performed on a BD Aria II cell sorter. All raw data files were processed using FlowJo software. For experiments where RNA was harvested, cells were sorted directly into 500μL of Qiagen Buffer RLT+, prior to RNA isolation (see below). For further propagation of live cells, cells were sorted directly into complete growth media. Experiments were performed in triplicate (at minimum) unless otherwise noted in figure legends.

RNA was harvested from cells using the RNeasy Plus Mini Kit (Qiagen), following the supplied protocol, and converted to cDNA using random oligomers and either Superscript III or Superscript IV reverse transcriptase (Invitrogen). qRT-PCR was performed on cDNA using the indicated TaqMan probes and analyzed on an Applied Biosystems StepOnePlus Real-Time PCR System. Relative quantification of mRNA abundance was performed using the 2-ΔΔCT method and ACTB or GAPDH as reference genes, where ΔCT = (CTtarget-CTreference) and 2-ΔΔCT = 2-(ΔCTsample-ΔCTcontrol). Note, in the cases of PAX2 and PAX8, for which no endogenous baseline expression was detected in ALL cells, baseline PAX5 (empty vector or vehicle) expression was used in calculating ΔCTcontrol. Experiments were performed in triplicate unless otherwise noted in figure legends.

At day 4 post-lentiviral transduction, 2×104 ZsGreen-positive cells of each PAX gene or control vector type were isolated by FACS and distributed into individual single wells of a 96-well plate. Beginning with normalized concentrations of 2×105 cells/mL (i.e., 100 μL total volume/well), these cells were further propagated in culture for a time course of 15–17 days. Culture density was assessed manually every 1–2 days during this time, using a hemocytometer. Additional media was added as needed prior to each counting in order to account for evaporation and to maintain ~100 μL volume in each well. For HEK293T cells, cell viability was assessed using an MTS assay (Cell Titer 96 One Solution, Promega), which produces a formazan product in the presence of phenazine methosulfate, which is present in metabolically active cells. Soluble formazan product is detectable at a 490nm absorbance maximum in PBS. Experiments were performed in triplicate (at minimum) unless otherwise noted in figure legends.

Cells were passaged one day prior to plating at a density of 2×105 cells/mL in 5 or 10mL of regular growth media with an added 80mM K-gluconate (Sigma, P1847) or CaCl2 (Sigma, C-3306) (unless otherwise noted). After indicated incubation times and depending on the experiment, RNA was either bulk harvested from treated cells or was harvested from live cells that were first sorted and collected by flow cytometry based on FSC-A and SCC-A measurements (indicated in figure legends). For pulse/chase in Fig 5E, 5F and 5G, cells were treated as indicated for 24 hours. 3×105 live cells were then sorted and returned to culture followed by removal of aliquots at indicated time points for harvesting of RNA (0h = 24 hour pulse, 0 hour chase). Experiments for all figures were performed in triplicate (at minimum) unless otherwise noted in figure legends.

4×106 Reh cells were electroporated with SMARTpool siRNA (i.e., 3 separate target siRNAs each) for PAX2, PAX5, NFAT5, or a non-targeting control pool (Dharmacon) using a BioRad GenePulser Xcell (Square wave, 210V, 15ms, 2x pulse, 0.1sec gap). Cells were suspended in 400μL Opti-MEM buffer containing 500nM siRNA that had been previously prepared and frozen at 20μM stock concentration in siRNA resuspension buffer (GE Healthcare). Cells were allowed to recover for 24 hours prior to harvest of protein lysates or treatment with hypertonic media (80mM K-Gluconate in RPMI with 10% FBS).