A branched epithelial ductal network is the architectural basis for the lung and collecting ducts of the kidney, as well as the salivary and mammary glands in mammals. Like other branched systems, the initial mammary branched pattern arises in embryonic development by crosstalk between the epithelium and the supporting mesenchyme (Wiseman and Werb, 2002; Parmar and Cunha, 2004). Unlike other branched systems, however, the mammary gland undergoes most of its branching at the onset of puberty, when circulating hormones, like progesterone and estrogen, begin systemic circulation (Woodward et al., 1998; Wiseman et al., 2003; Sternlicht et al., 2006; Fata et al., 2007; Khokha and Werb, 2011; McNally and Martin, 2011). In the mammary gland, epithelial ducts are the structural components that accomplish the function of conducting milk from the glandular component, called lobulo-alveolar units, to the nipple where it is consumed by a suckling newborn. The ductal portions of the mammary gland are composed of two layers of epithelium, an internal layer of luminal epithelium, and an external layer of contractile myoepithelium sheathed in a thin layer of basement membrane. The maintenance of this architectural form allows for apical basal cellular polarity, which is essential for proper luminal secretions and the primary function of the mammary gland (Adriance et al., 2005). These epithelial components are embedded in a complex stroma, which plays a considerable role in the specification and maintenance of this highly ordered branched structure (Kratochwil, 1969).
While the generation of this branched pattern occurs in a matter of weeks in the mouse, it is maintained over the lifetime of the organism. Therefore, uncovering the factors that contribute to the maintenance of branched architecture is equally important to understanding the developmental cues that result in the form and function of the mammary gland. It should be noted that although a branched architecture is maintained, morphological changes in lobulo-alveolar structures do occur with every reproductive cycle (Fata et al., 2001). Three-dimensional (3D) cultures of normal mammary epithelial cells using a laminin-rich ECM (lrECM) have been extremely valuable in determining the functional architecture and polarity of mammary epithelium (Bissell et al., 2003; Nelson and Bissell, 2005, 2006; Tanner et al., 2012). Continued research into the intricacies of branching morphogenesis in the mammary gland has highlighted the importance of extracellular factors such as hormones, extracellular matrix, mesenchyme, and growth factors, as well as numerous essential intracellular pathways, in the development of a branched structure (Simian et al., 2001; Sternlicht et al., 2005; Fata et al., 2007; Ewald et al., 2008; Andersen et al., 2011; Pasic et al., 2011; Basham et al., 2013). This heavy focus on better understanding branching morphogenesis in the mammary gland has provided seminal findings in the field of ductal development, as the mammary gland continues to provide researchers with a malleable model to investigate many aspects of biology.
Sodium Hydrogen Exchangers (NHEs) are a family of integral membrane proteins that catalyze the electroneutral exchange of extracellular Na+ for intracellular H+. The sodium hydrogen exchanger type 1 (NHE1) is a ubiquitously expressed member of this family that functions as a master regulator of pHi (Jang et al., 2006; Steffan et al., 2009). Currently, there are 9 subtypes of NHEs identified, with expression profiles following tissue- and membrane-specific patterns (Putney et al., 2002; Adriance et al., 2005; Slepkov et al., 2007; Casey et al., 2010). NHE1 has been found to have roles in cellular proliferation (Denker et al., 2000), migration (Denker and Barber, 2002; Stock et al., 2005, 2008; Stock and Schwab, 2006; Koliakos et al., 2008; Yang et al., 2010, 2011; Martin et al., 2011), intracellular acidosis-induced apoptosis (Lagadic-Gossmann et al., 2004; Pedersen, 2006; Wang et al., 2008; Casey et al., 2010), and critical developmental processes (Patel and Barber, 2005; Zhou and Baltz, 2013). Activation of this exchanger occurs by phosphorylation (Bianchini et al., 1997; Tominaga and Barber, 1998; Moor and Fliegel, 1999; Takahashi et al., 1999; Yan et al., 2001; Kintner et al., 2005; Luo et al., 2007; Malo et al., 2007; Meima et al., 2009), intracellular acidosis (Lagadic-Gossmann et al., 2004; Casey et al., 2010), or osmotic stress (Holt et al., 2006). Additionally, NHE1 has been shown to become activated as a result of growth factor stimulation (Moolenaar et al., 1983; Grinstein et al., 1988; Strazzabosco et al., 1995) and integrin activation (Tominaga and Barber, 1998). Previously, we have shown that its function is also necessary for mouse mammary branching morphogenesis, and that its inhibition alters growth factor–induced signaling (Jenkins et al., 2012).
In an effort to elucidate NHE1 function as is relates to the maintenance of organized tissue architecture, we first generated an established mammary branched structure using a previously described organotypic 3D-culture model. We then asked whether NHE1 was necessary to maintain tissue architecture in this model by exposing the tissue to the NHE1-specific inhibitor N-Methyl-N-isobutyl Amiloride (MIA). In doing so, we found that NHE1 inhibition resulted in a rapid loss of tissue organization and myoepithelial organization that was not accompanied by cell death or altered proliferation in our system. Additionally, NHE1 inhibition acidified intracellular pH (pHi) in end bud portions of branched tissue and altered actin localization in that region. Finally, we provide evidence that functional NHE1 is necessary for regulating E-cadherin in mammary epithelium. Our findings suggest NHE1 has a novel role in the maintenance of tissue architecture.
NHE1 Function is Required for the Maintenance of Mammary Tissue Branches
A fully established branched mammary tissue was first developed over the course of 4 days in a 3D tissue culture in order to ask whether NHE1 function was required to maintain this architecture (Jenkins et al., 2012) (Fig. 1A). After branches were established (Day 4) (Fig. 1A), NHE1 function was inhibited with 5(N-Methyl)-N-Isobutyl Amiloride (MIA) (50 μM) in the presence of growth factor TGFα (18 μM) (Fig. 1B) and tissue architecture was followed by time-lapse photography for the next 4 days (Day 4 to Day 8). MIA concentrations less than 50 μM did not overtly affect tissue architecture in the majority of structures (data not shown). In the absence of NHE1 inhibition, branches were maintained in both structure and number over the course of 4 days (Fig. 1B, top panels). In contrast, inhibition of NHE1 with MIA (50 μM) led to a rapid and drastic disruption of the branched architecture (Fig. 1B, bottom panels). Disruption was noted as early as 24 hr and involved the “fusing” of existing branches such that after 4 days only a mass of tissue existed, often containing few if any branches (Fig. 1B, compare bottom panels with top panels).
When compared to control tissue at Day 8 (4 days after MIA addition), inhibition of NHE1 function led to a greater than 65% reduction (100 ± 5.16 vs. 33.5 ± 10.59, control [adjusted to 100%] vs. MIA treated; P < 0.01) in the number of structures that exhibited at least 3 or more branches (Fig. 1C). Importantly, in tissue where NHE1 inhibition induced loss of architecture, TGFα-induced branching could be reinitiated after MIA was washed out of the culture system (Fig. 1D; and see Supp. Movie S1, which is available online). These findings indicate that maintenance of a branched mammary tissue requires NHE1 function and loss of NHE1 function induces a rapid non-permanent loss of tissue architecture.
NHE1 Inhibition Acidifies Intracellular pH in Mammary Tissue Endbuds
NHE1 is the key regulator of intracellular pH (pHi) in epithelial cells (Lagadic-Gossmann et al., 2004; Cardone et al., 2005; Casey et al., 2010). The exchanger performs this function by exchanging intracellular H+ for extracellular Na+ upon activation. Previously, we have shown that NHE1 inhibition with 10 μM of MIA disrupts branching morphogenesis and blocks growth factor–induced alkalization (Jenkins et al., 2012). Here we investigated the effect of NHE1 inhibition on pHi of branched structures 30 min after NHE1 inhibition by MIA (50 μM; Fig. 2). Using the ratiometric pH indicator BCECF-AM and our previous methods for detecting intracellular pH in mammary epithelial cells (Jenkins et al., 2012), we found that pHi in end buds of branched structures was significantly lower when NHE1 was inhibited (6.85 ± 0.02, N = 34 endbuds counted) compared to when NHE1 remained functional (6.97 ± 0.01; N = 19; P < 0.0001). Therefore, in 3D branched mammary tissue, the addition of MIA, which inhibits NHE1 function, rapidly induces intracellular pH acidification that can be effectively measured with the pH indicator BCECF-AM and ratiometric imaging.
Loss of Tissue Architecture Does Not Correspond to Increased Proliferation or Cell Death
NHE1 function has been shown to have a permissive role in both cellular proliferation (Pouyssegur et al., 1984) and acidosis-induced apoptosis (Li and Eastman, 1995; Lang et al., 2000). To characterize whether the loss of MIA-induced tissue architecture was a consequence of altered proliferation or cell death, we analyzed cell proliferation by phospho-histone H3 staining and cell death (necrosis, apoptosis, or autophagy) by Propidium Iodide (PI) exclusion, respectively (Fig. 3). An analysis of proliferation at Day 6 (0.11 ± 0.03% vs. 0.03 ± 0.04%, control vs. MIA-treated; P = 0.07) and Day 8 (0.57 ± 0.11 vs. 0.72 ± 0.13, control vs. MIA-treated; P = 0.7) indicated that any observed changes in tissue architecture were not associated with any significant differences in proliferation between control and MIA-treated samples (Fig. 3A). To investigate cell death, we stained live organoids at Day 8 with Propidium iodide (PI) to fluorescently label dead cells in which the plasma membrane had become compromised. Cell death analysis revealed no significant differences between control tissue (0.05 ± 0.006) and tissue treated with 50 μM MIA (0.04 + 0.005; Fig. 3B). We treated live tissue with 100 μM of MIA as a positive control for cell death, which induced approximately 100% of the cells to die after 4 days. These results show that inhibition of NHE1 function by MIA results in loss of mammary tissue architecture that is not associated with any observed changes in proliferation or cell death on day 6 or day 8.
Disruption of Mammary Tissue Architecture Within 24 Hours of NHE1 Inhibition
We noticed that a rapid loss of mammary tissue architecture occurred in the first 24 hr following NHE1 inhibition by MIA (50 μM; Fig. 4A). This loss of tissue architecture could also be induced by another NHE1-specific inhibitor ethylisopropylamiloride (EIPA) at the same concentration as MIA (Fig. 4B). To further explore this time window, we performed live-video microscopy of branched structures for the first 24 hr after NHE1 inhibition by MIA. An analysis of the resultant movies reveals that, when mammary tissue is inhibited in NHE1 function, it loses its architecture by a process of branch fusion, in which the end buds of the structure merge first, followed by the stalks (Fig. 4A and Supp. Movies S2 and S3). This process of branch fusion was rarely evident in mammary tissue not inhibited in NHE1 function (Fig. 1A; compare top with bottom panels). Moreover, propidium iodide staining of live tissue on day 5 indicated that this process was not accompanied by any significant pronounced necrosis, autophagy, or apoptosis (Fig. 4C). It should be noted that branch fusion could occur with the addition of MIA (50 μM) alone or when it was co-administered with TGFα (10 μM).
Tissue Polarity of Myoepithelial Cells is Disrupted When NHE1 Function is Inhibited
The mouse mammary gland has a distinct cellular organization pattern, with keratin 14 (K14) expressing myoepithelial cells surrounding keratin 8 (K8) expressing epithelial cells on the stalks of the branch, while K8-positive and -double positive (K8 and K14) cells are found at the tips of the end buds (Fig. 5A, left). Using immunocytochemistry for these keratins, we determined if this organizational pattern was distorted when the loss of architecture occurred after NHE1 inhibition. Within 24 hr of NHE1 inhibition (Day 5), the continuous sheet of K14-postive myoepithelial cells seen in the stalk areas did not exist anymore (Fig. 5A). Instead, K14-positive cells became discontinuous throughout the entire surface of the tissue (Fig. 5A). We determined that the changes in K14-positive cells were likely not a consequence of ectopic expression since the amount of K14 protein was equal between control tissue and MIA-treated tissue (Fig. 5B). To further analyze the K14-phenoytpe, we performed optical sectioning of K14-immunostained tissue by confocal microscopy (Fig. 5C). This analysis revealed that in mammary tissue inhibited in NHE1 function, myoepithelial cells could be found on the surface as well as deep within the altered tissue as compared to control tissue where myoepithelial cells are only found on the external surface (Fig. 5C). By examining the apical marker ZO-1 in relationship to myoepithelial cells (K14), it becomes apparent that myoepithelial cells found deep within the tissue can be found in areas absent of Z0-1 and in many areas where ZO-1 is prominent. Although this latter finding indicates myoepithelial polarity can be reversed when NHE1 is inhibited, there are some instances in control tissue where the marker K14 can be found in apical areas (ZO-1 positive). These K14+/ZO-1+ areas may likely represent cells found to be double positive for both K8 and K14 (Fata et al., 2007). Together these changes indicated that altered tissue organization was accompanied in areas where abnormal tissue polarity of myoepithelial cells is evident, and all occurring within 24 hr after NHE1 inhibition.
Inhibition of NHE1 Function Leads to a Loss of NHE1 Basal Polarity
In epithelial cells, NHE1 can be found to exhibit prominent basal-lateral polarity with a marked absence from the apical surface (Damkier et al., 2009). As expected, in mammary tissue freshly isolated from the mouse, immunostaining revealed both myoepithelial and epithelial cells express NHE1 and that in areas containing lumens, a basal-lateral polarity is evident with lower or absent expression on the apical side (Fig. 6A). In mammary tissue that had undergone branching in our assay, normal NHE1 expression and basal-lateral polarity was still maintained throughout the branched structure (Fig. 5B, top panels). However, in mammary tissue inhibited in NHE1 function, the basal polarity of NHE1 became disrupted and lost in a number of areas. Under these conditions, where NHE1 function was inhibited, basal polarity of NHE1 appeared to either reverse or become lost, such that NHE1 became absent on the basal side and evident on the apical side (Fig. 5B, bottom panels). When we quantified this change of polarity, we found that NHE1 inhibition led to an 8.5-fold increase of cells where basal polarity was absent on the periphery of disrupted structures compared with branched structures where NHE1 function was not perturbed (Fig. 6B, P < 0.0005). Therefore, the rapid loss of tissue architecture caused by NHE1 inhibition is accompanied by a significant change of NHE1 basal-polarity in a number of cells.
F-actin and E-cadherin Are Altered by NHE1 Inhibition
Since we observed rapid fusing of branches during the loss of architecture, we hypothesized that NHE1 inhibition could alter the cytoskeleton and/or cell–cell adhesion molecules necessary for maintaining epithelial structures. Filamentous actin (F-actin) is a structural cytoskeleton protein known to have an important role in maintaining cellular and tissue structure (Yamazaki et al., 2007; Tang and Brieher, 2012) and is known to be affected by changes in pHi (Lagarrigue et al., 2003). In an effort to determine the pattern of F-actin organization in both the day-5 branched structure (−MIA) and the day-5 disrupted structure (+MIA), we stained both conditions with phalloidin. We found that branched tissue had thick rings of circumferential actin on the apical side of the end bud lumens (Fig. 7A, top panels), consistent with what has been observed in other polarized epithelial systems (Yonemura et al., 1995; Smutny et al., 2010). In contrast, in tissue where the architecture was disrupted by NHE1 inhibition, these thick highly organized bands were not observed at any z-plane (Fig. 7A, bottom panels). No other differences were apparent when examining F-actin by phalloidin labeling. To investigate cell adhesion molecules, we cultured mammary primary epithelial cells in 2D, exposed them to MIA for 24 hr, and then immunostained for E-cadherin. When confocal florescent images were then captured under identical conditions and identical z-planes, we found NHE1 inhibition associated with a decrease in E-cadherin expression (Fig. 7B). Lower levels of E-cadherin expression were also noted in tissue cultured in 3D that had become disrupted due to NHE1 inhibition (Fig. 7C). In 3D cultures, this loss of E-cadherin expression was most prominent deep within the tissue mass and, in both 2D and 3D mammary epithelial cultures, E-cadherin loss could be found within 24 hr of inhibiting NHE1 with MIA. We had noted that organoids were able to reorganize themselves back into branched structures once NHE1 inhibition was removed from 3D cultures. We, therefore, hypothesized that E-cadherin expression would need to be restored to normal levels for this to occur. To test this hypothesis, we isolated protein from organoids that were cultured in 2D and probed for E-cadherin expression by immunoblot. We found that NHE1 inhibition for 24 hr lowered E-cadherin expression and that this effect could be rescued once the NHE1 inhibitor was removed from culture for 72 hr (Fig. 7D; compare lanes 2–4). These results provide strong evidence that, in mammary tissue, NHE1 function is necessary to maintain both normal F-actin cytoskeleton arrangement and E-cadherin expression.
Branching morphogenesis is a highly ordered and efficient process that generates a ductal network critical to the function of a number of biological structures, including the mammary gland. Perhaps as interesting as how a branched structure is generated is how these structures are maintained in form and function over the lifetime of the organism. Previously, we found that NHE1 inhibition with 10 μM MIA dramatically disrupted growth factor–induced branching morphogenesis, indicating this hydrogen exchanger is integral to this developmental process. These original findings also concluded that 48 hr after growth factor stimulation, inhibition of NHE1 function with 10 μM MIA had no effect on branching, suggesting that the early signaling events require NHE1 activity. Here we demonstrated that NHE1 is also necessary for the maintenance of branched tissue architecture and that its inhibition with a higher concentration of MIA (50 μM) resulted in a rapid but reversible loss of tissue organization. Together, these findings point to a threshold effect, where partial NHE1 inhibition can affect the early events of branching morphogenesis without affecting an established tissue architecture, while a more complete NHE1 inhibition can affect tissue architecture. Along this spectrum, complete inhibition of NHE1 function with 100 μM of MIA leads to rapid cell death throughout the entirety of the tissue (Jenkins et al., 2012).
Establishing a functional branched structure requires a dynamically regulated interaction between the stroma and epithelium, as well as between the epithelium and the basement membrane (Bissell et al., 2002, 2003; Bissell and Bilder, 2003; Parmar and Cunha, 2004; Adriance et al., 2005). These interactions provide vital contextual cues that regulate processes such as cellular polarity, which is critical for vectoral milk production in mammary epithelium (Barcellos-Hoff et al., 1989), and proliferation, which is controlled within specific windows during branching morphogenesis (Fata et al., 2007; Ewald et al., 2008). Additionally cell migration, a process that requires both regulated cell adhesion and modification of the ECM, has been shown to play a pivotal role in driving mammary branching morphogenesis (Ewald et al., 2008) and in generating epithelial tubes (Mori et al., 2009). It stands to reason that the regulated pathways necessary to establish mammary tissue architecture are also necessary to maintain it. The importance of these pathways is no better illustrated then when one considers cancer, where functional tissue disregards normal polarity and growth regulating pathways and begins to proliferate in a disorganized and dysfunctional manner.
Na+/H+ exchanger function has been shown to be involved in establishing cellular polarity (Patel and Barber, 2005) and in creating favorable micro-environments for ECM interaction and actin stabilization (Bernstein et al., 2000), as well as actin filament organization downstream of growth factor stimulation (Meima et al., 2009). Importantly, actin regulation plays a critical role in maintaining epithelial cell interaction by E-cadherin (Yamazaki et al., 2007; Tang and Brieher, 2012), and has been shown to be dynamically regulated at cell–cell contacts (Kovacs et al., 2011; Yamada et al., 2012). Here, we have shown the perturbing NHE1 function altered both actin organization and E-cadherin expression resulting in a reversible loss of branched tissue architecture. This decrease in E-cadherin is similar to what has been reported during mammary branching morphogenesis, specifically when the epithelium is rearranging during ductal elongation (Ewald et al., 2008). Taken together, a possible mechanism for our observations arises. NHE1, by its ability to maintain pH homeostasis, contributes to the stabilization and localization of actin to the zona adherins of polarized epithelium. When NHE1 function is altered, dynamic actin localization in the apical perijunctional region is disrupted and E-cadherin protein is decreased, thus making tissue architecture unstable. Our findings suggest a novel homeostatic regulatory pathway that may involve NHE1 as a possible upstream regulator of E-cadherin expression and ultimately tissue stability.
The loss of mammary ductal architecture is clearly evident in the formation of tumors, which develop from established ducts by disregarding the influence of normal polarity pathways. Therefore, understanding the factors that maintain mammary tissue architecture not only feeds our understanding of mammary gland development and function, but also provides information that impacts how polar tissue becomes apolar during tumor development. Both intracellular and extracellular pH is known to be deregulated in cancer and is thought to play a major role in the progression of the disease (Cardone et al., 2005; Stock and Schwab, 2009). Here, we have found that deregulation of intracellular pH leads to loss of tissue architecture analogous to cancer progression. A number of studies have shown that cellular context and tissue architecture greatly influence cancer phenotypes despite considerable genetic aberration (Mintz and Illmensee, 1975; Wang et al., 2002; Kirshner et al., 2003). Our results suggest a novel role for NHE1 and pH homeostasis in maintaining normal tissue structure and function independent of direct genetic manipulation. This role should not be underestimated when considering that the maintenance of tissue architecture is absolutely necessary for mammary function and that loss of this architecture is a hallmark of breast cancer.
Mammary Tissue (Organoid) Isolation
Mouse mammary tissue was isolated and cultured as previously described (Fata et al., 2007; Jenkins et al., 2012). Briefly, the fourth inguinal mammary glands were isolated from virgin Balb/C mice 14–30 weeks of age, diced with two standard razor blades, and placed in 10 mL of a trypsin/collagenase mix (0.2% trypsin/0.2% collagenase, 5% fetal calf serum (FCS), 1,000 U/ml Penicillin/Streptomycin) for 25 min gently shaking at 37°C. Following digestion, the suspension was centrifuged for 5 min at 70g. The pellet was washed twice in 7 mL of DMEM/F12 from CellGro and was then resuspended in 4 mL of DMEM/F12 with DNase (2U/μL) gently shaking at room temperature for 5 min. The larger pieces of mammary gland (mammary organoids) were then separated by differential centrifugation, i.e., pulse spins to 70g at least three times with the supernatant being discarded every time. The final pellet was used for three-dimensional (3D) mammary tissue cultures.
Three-Dimensional (3D) Branching and Loss of Architecture Assay
Three-dimensional culturing of mammary tissue in matrigel was performed as previously described (Fata et al., 2007; Jenkins et al., 2012). A 40-μL supporting layer of matrigel (BD Sciences, San Jose, CA) was first added to each well in a 96-well format and allowed to solidify in the incubator (37°C and 5% CO2) for 30 min. Isolated mammary organoids were suspended in 100% matrigel plated at 40 μl/well and at a density of 50 organoids/well. The organoid containing layer was allowed to solidify for 30 min before the addition of 100 μl of basal media, also called mammary epithelial cell (MEC) media (DMEF/F12 with 1% insulin, transferrin, selenium, and 1% penicillin/streptomycin). After 24 hr, organoids were stimulated with TGFα (18 nM). This was considered day 0. Media was changed to basal media after 48 hr (day 2). Branching was considered complete on day 4 at which time organoids were re-stimulated with TGFα (18 nM) in either in the presence of 5(N-Methyl)-N-Isobutyl Amiloride (MIA; Sigma-Aldrich, St. Louis, MO; 50 μM) or in the absence of MIA. Ethylisopropylamiloride (EIPA; Sigma-Aldrich), another NHE1-specific inhibitor, was also examined in this assay. Media was again changed to basal (MEC) media alone on day 6. The assay was considered complete 2 days later on day 8 when all remnants of organoid branches were no longer apparent following NHE1 inhibition. A structure was scored as branched if it had three or more distinct branches (end buds and stalks) protruding in separate directions (Fata et al., 2007). See Figure 8 for a schematic representation of this assay. To capture the loss of architecture, live video microscope was conducted as previously described (Jenkins et al., 2012). Briefly, cultures were kept in a fully enclosed inverted Microscope (Zeiss Axio Observer, Thornwood, NY) and imaged by brightfield microscopy every 1 or 24 hr.
Two-Dimensional (2D) Culture of Organoids
Mammary organoids were isolated as described above and cultured in MEC media supplemented with 2% fetal calf serum (FCS) at a density of 100 organoids/well in a 48-well plate. Organoids were allowed to adhere and spread out for 48 hr before being serum starved for 24 hr in MEC media supplemented with 0.5% FCS. The tissue residues were not removed directly, however many washed away in subsequent media washes. All cultures were serum starved before experimentation. Subsequent experimental materials (growth factors and/or inhibitors) were suspended in MEC media supplemented with 0.5% FCS. For recovery from NHE1 inhibition, organoids were cultured in MEC media containing 2% FCS once the NHE1 inhibitor was removed.
Cell Death Assay
Cell death in organoids was assessed as described previously (Jenkins 2012). At day 5 and at 8, live tissue was stained with propidium iodide (PI) (100 μM) to detect cells in which the plasma membranes had become compromised by necrosis or apoptosis. Tissue was then imaged by brightfield and fluorescent microscopy. Cells positive for PI staining were scored as dead. Quantification was performed by dividing the number of dead cells by the area of the tissue imaged. Area was measured using ImageJ.
Immunostaining for NHE1, Phospho-Histone H3, and E-Cadherin
Mammary organoids cultured in 3D were fixed with 4% paraformaldehyde at 37°C for 5 min. The cultures were then washed two times (5 min each) with phosphate buffered saline (PBS) before being permeabilized with PBST (PBS+ 0.5% Triton X-100) for 10 min. Blocking was performed with blocking buffer (10% horse serum, 2% bovine serum albumin, and 0.5% Triton X-100 in PBS) for 1 hr. Primary antibodies against phospho-histone H3 (1:100)(Ser10, Millipore, Billerica, MA), NHE1 (1:50) (clone 2F5: WH0006548M1; Sigma-Aldrich) (H-160:sc 28758; Santa Cruz Biotechnologies, Santa Cruz, CA.), or E-cadherin (clone EP700Y; Millipore) were suspended in blocking buffer and incubated overnight at 4°C. After incubation with primary antibody, samples were washed (2 times with PBS for 5 min each, then 1 time with PBST for 5 min) and incubated with a species appropriate Alexa Fluor conjugated secondary antibody (Invitrogen, Carlsbad, CA; 1:1,000) in blocking buffer for 1–3 hr shaking at room temperature. After another wash series, samples were stained with 4′,6-diamidino-2-phenylindole (DAPI) or propidium iodide (PI) for 5 min and imaged. Quantification of phospho-histone H3 staining was performed by capturing optical sections of organoids and dividing the number of positive phospho-histone H3 cells by the total number of cells found in the structure by PI staining. Quantification of NHE1 polarity was accomplished by capturing optical sections of organoids. Cells on the periphery from the center slice of each structure were scored for polarity.
Immunostaining for Keratin 8 (K8), Keratin 14 (K14), and Z0-1
3D cultures were fixed with methanol/acetone 1:1 for 20 min at −20°C. They were then rinsed twice with PBS for 5 min in the 96-well culture plates. The matrigel was then removed from the well and partially air-dried onto an 8-chamber slide (BD Sciences). Samples were then blocked as described above. Primary antibodies against keratin-14 (K14) (clone LL002, Novocastra, UK) (1:100), keratin-8 (K8) (Troma-I-c, DSHB, Iowa City, IA) (1:100), or zonulae occludins-1 (ZO-1; clone 1, BD Biosciences) were suspended in blocking buffer and incubated overnight at 4°C. Secondary antibody staining was performed as described above for NHE1 and phospho-histone H3 staining.
BCECF-AM Staining for Detection of Intracellular pH (pHi)
BCECF-AM was used exactly as described previously (Jenkins et al., 2012). Briefly, day-4 branched organoids were stimulated with TGFα (18 nM) ± MIA (50 μM) in the presence or absence of NHE1 inhibition by MIA for 30 min before pHi was measured. The structures were then incubated with BCECF for 5 min and imaged by fluorescent microscopy using a filter set designed specifically for BCECF-AM (Carl Zeiss). BECEF-AM-loaded organoids were then alternatively excited with 440 and 490 nm light, recording emission at 535 nm. The light source was a mercury bulb. An image (1,390×1,390) was captured at each excitation wavelength. Ratiometric analysis, i.e., pixel by pixel division of the 440- and 490-nm images, and a standard curve of pHi were then used to determine the pHi as described previously (Jenkins et al., 2012). The ratio image was then mapped to a discrete 16-bit color map so that pH values could be visualized.
Actin Staining With Phalloidin
Actin organization was visualized by staining with fluorescent conjugated phalloidin (Alexa Fluor® 594 phalloidin from Invitrogen). Organoids were cultured in a 96-well format and stimulated to branch (day 0), then re-stimulated (day 4) with TGFα in the presence or absence of NHE1 inhibition by MIA (50 μM). Organoids were fixed with the Actin Visualization Biochem Kit from Cytoskeleton (Cat. BK005) on day 5. Fixation and staining were performed in the well according to the manufacturer's specifications. After staining was complete, the samples were removed from the well and lightly air-dried to a glass slide to be imaged by confocal microscopy.
Immunoblotting was performed as previously described (Jenkins et al., 2012). All primary antibodies were suspended in Tris buffered Saline (TBS) containing 5% BSA and 0.05% Tween-20. Rabbit anti-E-cadherin (Cell Signaling, Danvers, MA; 3195) (1:1,000) was used to detect E-cadherin. Mouse monoclonal anti-keratin-14 (K14) (clone LL002, Novocastra) (1:100) was used to detect keratin 14. β-Actin was detected with rabbit polyclonal anti-β-Actin (Cell Signaling, 4967) (1:2,000). All samples were briefly sonicated on ice prior to loading.
Statistical significance was determined using the Student's t-test function in GraphPad Prism 5 for Windows (GraphPad Software, San Diego, CA).