extfieldsofstudy: Biology Medicine

text:
An Investigation into the Immunomodulatory Activities of Sutherlandia frutescens in Healthy Mice

Sutherlandia frutescens is a medicinal plant that has been traditionally used in southern Africa for cancers, infections, and inflammatory conditions. We recently published experiments demonstrating that an aqueous extract of S. frutescens possessed potent immune-stimulatory activity. This work was carried out with murine macrophages, an immune cell type that plays a pivotal role in host defense from infection and in shaping host inflammatory and immune responses. Here, we conducted a series of follow-up experiments to explore the impact of consuming S. frutescens on host response to bacterial challenge using healthy mice. We found that feeding mice a diet containing S. frutescens failed to significantly alter host response to systemic infection by either a gram-positive or gram-negative bacterium (i.e., L. monocytogenes and E. coli, respectively). In contrast to the in vitro observations, we found no evidence that S. frutescens consumption stimulated in vivo inflammatory responses; instead, consumption of S. frutescens tended to diminish in vivo inflammatory responses. Several possible reasons for this are discussed.

Introduction
Recently we used a murine macrophage cell line and primary murine macrophages to demonstrate that extracts of S. frutescens have potent immune-stimulatory activities. [1] Specifically, short-term treatment with a crude aqueous extract of S. frutescens or with a polysaccharideenriched fraction of S. frutescens increased macrophage production of reactive oxygen species (ROS) and nitric oxide (NO). ROS are a group of highly reactive free radical and non-radical molecules, including superoxide (O 2 • -), hydrogen peroxide (H 2 O 2 ), hydroxyl radicals (OH•), hypochloric acid (HOCl), and peroxynitrite (ONOO-). [2] NO has been recognized as a versatile mediator of immune responses. [3] Both ROS and NO play critical roles in host defense to infection. [4][5][6] Additionally, the aqueous extract and polysaccharides-enriched fractions of S. frutescens activated NF-κB signaling in murine macrophages resulting in an increased period, mice were randomly assigned into three dietary treatment groups: 0, 0.25 or 1.0% (wt/ wt) ground S. frutescens leaf powder in AIN-93G basal rodent diet. [13] Experimental diets The botanical-containing diets were made using the ground powder of the vegetative parts of Sutherlandia frutescens purchased from Big Tree Nutraceutical (Fish Hoek, South Africa) and authenticated as described previously. [1] According to previous reports, the most commonly used dose of S. frutescens in humans is 2.5 g dry materials (leaf) per day. [14] For a 70 kg adult human, this dose of S. frutescens intake would be equivalent to 35.7 mg/kg per day. Therefore, we used 36 mg/kg per day to calculate the human equivalent dose for experimental diets in our mouse feeding studies. The human equivalent dose was calculated using a formula that adjusts for differences in metabolic body size and is described in detail elsewhere. [15,16] For mice this dose was estimated to be 411 mg/kg body weight per day. The average body weight of our healthy adult mice was~25 g, therefore we estimated that each mouse would need to consume 11 mg S. frutescens to achieve the dosage of 411 mg/kg BW per day. Food consumption for these mice was expected to be 4-5 g/d based on our previous experience and reports in the literature, thus to deliver~11 mg of S. frutescens per day, the diets should contain approximately 0.25% (g/100 g) of the ground S. frutescens material. We also prepared a diet containing four times this dosage (i.e., 1.0%) of S. frutescens to account for unexpected and unknown differences between mice and humans relative to the uptake and metabolism of putative bioactive compounds in S. frutescens. All diets were nutritionally complete [13] (i.e., AIN-93G diets) and identical, except for the presence of S. frutescens and the small amount of cornstarch that was removed to account for the addition of this botanical ( Table 1).

In vivo bacterial challenges
To assess the impact of dietary S. frutescens on host-resistance from infection by L. monocytogenes we used healthy male and female BALB/c. This strain of mice is known to be susceptible to infection from L. monocytogenes. [17] After 4 weeks of consuming the experimental diets, all mice received an intravenous injection with~10 4 cfu of L. monocytogenes (EGD strain) in 0.2 mL of sterile, endotoxin-free phosphate-buffered saline.
To investigate the impact of dietary S. frutescens on host response to an E. coli infection, male and female C57BL/6 mice were used. After consuming one of the experimental diets for 4 weeks, mice were injected intraperitoneally with~10 8 colony-forming units (cfu) of E. coli (K-12 strain) in 1 mL of sterile PBS.
Following bacterial challenge, mice were monitored twice daily for clinical symptoms of illness using a standard scoring sheet designed for monitoring and recording pain/discomfort in rodents and approved by the Animal Care and Use Committee of the University of Missouri. Any mice showing signs of pain or distress (e.g., ruffled fur, shallow breathing, hunched back) were immediately and humanely euthanized. As expected, no mice in our experiments experienced clinical symptoms requiring premature termination, however, we did record three premature deaths during the E. coli challenge experiments (i.e., female mice in CONTROL and 0.25% SF groups and a male mouse in the 1.0% SF group) and one unexpected death during the L. monocytogenes experiments (i.e., a female mouse in the 0.25% SF group).

Sickness behavior
The impact of S. frutescens on spontaneous locomotor activity before and during an infection was measured by mini-emitters as described by Spiers et al. [18] Representative mice (n = 4 of each sex) within each diet treatment group were lightly anesthetized and mini-emitters were surgical implanted subcutaneously along the spine just below the scapulas. Following a 4-day post-surgery recovery period, activity data were recorded every 5 seconds with accumulated events for every 20 min period for a 3-day period prior to and 3 days or 36 hours following the challenge with L. monocytogenes or E. coli, respectively, following the bacterial challenge.

Measurement of host clearance of the bacterial challenge
We determined the number of L. monocytogenes present in the spleen and liver of mice 3 days following the challenge. This timeframe best reflects the capacity of the innate immune system to control the cell-to-cell spread and proliferation of this intracellular pathogen. [19,20] Therefore, 3 days following the challenge with L. monocytogenes mice were humanely killed and the liver and spleen from each mouse was aseptically collected and weighed. One hundred milligram portions of each tissue were transferred to 1.5 mL sterile centrifuge tubes containing 0.2 mL of sterile PBS and 100 mg of 0.5mm zirconium oxide beads. Tubes were kept on ice until processed for bacterial load determination as described below.
The bacterial load in the liver and spleen was determined by a fluorescence-based microplate assay. [21] The assay is based on the reduction of the non-fluorescent dye resazurin to a fluorescent product, resorufin, by bacterial enzyme activity. Liver and spleen samples were collected and then homogenized using a Bullet Blender TM (Next Advance, Inc., New York, United States of America) according to the manufacturer's instructions and described elsewhere. [22] One hundred microlitersof the homogenates were diluted to 1.0 mL (10-fold) with PBS containing 0.2% saponin, then 7 μL of these diluted homogenates were added to wells of a black 384 well plate in triplicate. Each well contained 50 μL of 0.01% resazurin in LB medium. Resorufin fluorescence was measured every 20 minutes for 10 hours. The time required to reach 8000 RFU (T 8K ) was determined for each sample and cfu/mg tissue calculated using the standard curve. A suspension containing 1x10 9 cfu/mL of L. monocytogenes or E. coli was prepared, and serial 10-fold dilutions of this suspension were done in ice-cold LB medium containing 0.01% resazurin to create standards in the range 1x10 2 to 1x10 8 cfu/mL. Fifty microliters of these standards were added in triplicate to a pre-chilled black 384 well plate and fluorescence (excitation, 530 nm; emission, 590 nm) was read every 20 min for 10 h using a fluorescence plate reader (BioTek Synergy HT). A standard curve was then constructed that relates the log of the cfu/well to the time required to reach a standard fluorescence of 8000 relative fluorescence units (RFU). The limit of detection for this assay was approximately 4x10 3 cfu/g tissue. For liver samples that were below the limit of detection in the fluorescence assay, cfu/mg tissue was calculated using a traditional microbiological plate count approach. For these liver samples only, 100 μL of the diluted homogenate was plated on TSAY (tryptic soy agar + yeast extract) plates. Plates were incubated overnight at 37°C and colonies counted manually the next day.

In vivo host inflammatory response
In order to assess the impact of S. frutescens consumption on host inflammatory response, BALB/c and C57BL/6 mice were fed with experimental diets as described previously, followed by an intraperitoneal injection with L. monocytogenes (EGD strain, 5 x 10 6 cfu/mouse) and LPS (from E. coli 026:B6, 20 μg/mouse), respectively. Mice were anesthetized 2 hr post-LPS injection or 24 hrs. after L. monocytogenes challenge and blood was drawn by cardiac puncture. After allowing the blood to clot on ice for 30 minutes, serum was separated by centrifugation (1000× g, 20 min.) and stored at -80°C for later analyses of inflammatory cytokines/chemokines as described below.

Isolation and stimulation of peritoneal macrophages
Thioglycolate-elicited peritoneal macrophages were isolated from healthy mice as described previously. [23] Crude (i.e., unfractionated) peritoneal cells were re-suspended in Dulbecco's Modified Eagle's Medium (DMEM) with 5% FBS, and seeded into a 96-well plate at a density of 1×10 5 cells/well. Cells were cultured in the incubator at 37°C with 5% CO 2 for 3 h. The nonadherent cells were removed by washing with PBS, leaving behind a monolayer of macrophages (typically > 95% purity).
We used a commercial synthetic diacylated lipoprotein (i.e., Pam2CSK, Invivogen, San Diego, CA, USA) to mimic L. monocytogenes infection for these ex vivo experiments, while LPS served to model E. coli infection. The production of inflammatory cytokines/chemokines by elicited cells was also evaluated. We conducted dose-response experiments to determine the optimal concentration for each of these inflammatory stimuli in our assay. The concentrations used (i.e., 5 ng of Pam2CSK or 100 ng of LPS per mL of culture medium) provided a robust, but sub-maximal stimulation. Primary macrophages were co-cultured with LPS or Pam2CSK for 24 h, and the cell culture medium was harvested, stored at -80°C for later cytokines and chemokines quantitation.

Statistical analyses
All data were analyzed by one-way analysis of variance (ANOVA) using SAS 9.3 software (SAS Institute Inc., Cary, NC, USA). The post-comparisons were performed using Tukey's multiple comparison tests. A p < 0.05 was considered to indicate statistical significance. Data are presented as means ± SEM, except on those occasions when unequal variances existed between independent trials or assays, then LSmeans ± pooled SEM are reported.

Host resistance to bacterial infection
After consuming diets with 0.25% or 1.0% S. frutescens for~4 weeks, the average body weight of female and male mice (both BALB/c and C57BL/6 strains) were not significantly different within sexes from that of control mice (see S1 and S2 Tables, respectively). Challenging mice with L. monocytogenes resulted in a loss of 5 to 10% of their body weight during the next few days, regardless of which experimental diet the mice were consuming. Similarly, consumption of S. frutescens had no discernable impact on the loss of body weight in mice following an E. coli challenge. Additionally, there were no significant differences across diet treatment groups in liver or spleen weight following either bacterial challenge.
The impact of consuming S. frutescens containing diets on host defense against infection from two commonly studied bacterial pathogens is shown in Figs 1 and 2. In contrast to our expectations, there was little indication that consumption of S. frutescens enhanced host clearance of either of these bacteria during the early stages post-challenge. We did however, note a trend (p < 0.08) for a modestly lower burden (~0.5 log 10 ) of L. monocytogenes in the livers of male BALB/c mice fed the 0.25% S. frutescens diets compared with the mice fed the control (0% SF) or 1% S. frutescens diets (Fig 1C). Consumption of S. frutescens appeared to have no noticeable impact on the burden of L. monocytogenes in the spleens of male mice, nor on bacterial burden in the livers and spleens of the female BALB/c mice (Fig 1A, 1B and 1D). Additionally, there were no statistically significant differences in E. coli burden in livers or spleens of female or male C57BL/6 mice fed either S. frutescens-containing diets compared to the control diet (Fig 2A-2D).

Impact of infection on spontaneous activity (i.e., sickness behavior)
We implanted mini-emitters in a separate group of mice to monitor the impact of S. frutescens consumption on overall wellbeing of the mice immediately prior to and during bacterial challenges. The spontaneous activity (i.e., random locomotion within the cage) of four individual female mice in each dietary treatment group before and after infection with either L. monocytogenes or E. coli is shown in Fig 3 and Fig 4, respectively). Not surprisingly, spontaneous locomotor activity dramatically declined following the infectious challenge with either bacterial pathogen. Dietary S. frutescens consumption appeared to have no discernable impact on this activity either before or during the infection. While data from female mice are shown, similar results were observed for male mice similarly challenged with L. monocytogenes or E. coli (see S1 and S2 Figs). After consuming diets for~4 wks, all mice received an intravenous injection with~10 4 cfu of L. monocytogenes (EGD strain) in 0.2 mL of sterile, endotoxin-free phosphate-buffered saline. Liver and spleen samples were harvested three days after challenge and colony-forming units (cfu) were determined by a fluorescence-based microplate assay of tissue homogenates. Each symbol represents the data from a single mouse with the median and intra-quartile range shown. In vivo production of cytokines and chemokines Systemic infection with L. monocytogenes in previously healthy BALB/c mice induces the production of numerous cytokines and chemokines quantified from blood samples collected 24 h post-challenge (Table 2). Monocyte chemotactic protein-1 (MCP-1, CCL2), interferon(IFN)-γ, and interleukin-6 (IL-6) were the dominant factors detected in the blood of L. monocytogenesinfected mice. We observed no significant differences in the measured cytokines and chemokines based on the experimental diet consumed prior to and 24 hours following the infectious challenge. To evaluate the impact of S. frutescens intake on the inflammatory response in healthy mice we challenged them with lipopolysaccharide (LPS) from E. coli. Mice received an intraperitoneal injection with 20 μg of LPS followed by blood collection 2 hours later. We observed that consumption of S. frutescens by healthy C57BL/6 mice tended (p < 0.09) to reduce by nearly  50% the production of tumor necrosis factor-alpha (TNF-α) in response to a LPS challenge ( Table 3). As can be seen in Table 3, however, we failed to detect any other diet-induced differences in the circulating cytokines, chemokines, or growth factors quantified. We also did not observe any significant impact of S. frutescens consumption on basal or background cytokine/ chemokine levels in mice injected with vehicle (i.e., PBS) alone (see S1 Table).

Ex vivo cytokines and chemokines production
Primary peritoneal macrophages isolated from mice fed S. frutescens produced significantly less IL-1α following stimulation with LPS compared to macrophages isolated from mice fed the control diet (S4 Table). In contrast, ex vivo production of other cytokines/chemokines by LPS-or Pam2CSK-stimulated macrophages were not significantly impacted by dietary S. frutescens (S4 and S5 Tables).

Discussion
With most botanicals for which immune-modulating activity is claimed the evidence of their activity is limited to actions on isolated immune cells (i.e., in vitro). Previously, we reported  that a hot-water extract made from the medicinal plant S. frutescens potently activated murine macrophages, resulting in increased production of ROS and NO. [1] Production of ROS and NO by innate immune cells are early and critical components of host defense against infection from bacterial pathogens. [4,5] We found that this immune-stimulating activity was associated with a polysaccharide-enriched fraction of this aqueous extract. Zhang et al. [24] found that pectin-type polysaccharides isolated from Lessertia frutescens (an alternative name for Sutherlandia frutescens) stimulated innate immune cells via complement fixation. The complement cascade is a part of the host's immune response that promotes host clearance of microbes and damaged cells. There exists a large body of research showing that numerous botanical polysaccharides possess in vitro immune-stimulatory activity. [9] In the present study, however, we investigated the potential benefits of unfractionated vegetative parts of this botanical in response to bacterial infection. To our knowledge this is the first investigation to examine in vivo and ex vivo host anti-bacterial activity associated with the consumption of S. frutescens.
In the present study, we found that feeding healthy mice diets containing S. frutescens had little impact on the host's immune response to either of the bacterial pathogens we tested. In light of the results from our prior in vitro experiments, we designed the present experiments to focus on the early or innate elements of the host's immune response to infection. Following the following challenge, following sera isolation, specific cytokines and chemokines concentrations were determined using a commercial multiplex beads-based assay system. Data shown are from 12 mice from each diet group (i.e., 6 mice per diet treatment group in each of two independent trials), with values representing the means ± SEM. Values for T-cell derived cytokines (i.e., IL-2, IL-4, IL-5, IL-7, IL-9, IL-13, IL-15, and IL-17) tended to be very low in concentration; were not significantly impacted by the diet treatment, thus were not reported. b The impact of the diet intervention/treatment was tested in SAS by ANOVA, using contrast with interaction between the two independent trials/experiments with the main effect p-value shown in the 4 th column.
c Abbreviations (refer to intravenous administration of L. monocytogenes, these gram-positive bacterium are rapidly taken up by phagocytes, primarily in the spleen and liver. [17] Intracellular pathogens, such as L. monocytogenes, successfully replicate within these phagocytes unless the generation of ROS and NOS are stimulated. [12] Whether and to what extent the host is able to prevent replication and spread of this intracellular pathogen during the early stages of infection (e.g., 3 days postchallenge) is highly dependent upon the production of a variety of host factors, including: TNF-α, [25] IFN-γ, [26] IL-6 [27] and MCP-1 [28]. Therefore, we expected to find evidence that this botanical modified the early production of these host factors and that mice fed diets containing S. frutescens would contain fewer of these bacteria. However, when we sampled the blood just 24 hours post-challenge with L. monocytogenes we found no indication that consumption of S. frutescens had altered the production of these key cytokines and chemokines. Consistent with this result was our failure to observe a significant difference in the number of L. monocytogenes bacterium in either the spleen or liver of mice fed diets containing S. frutescens. Similar to our results examining in vivo host response to a L. monocytogenes infection, we found that dietary S. frutescens failed to modulate the host response to a challenge with a common gram-negative organism, E. coli. Yet, we did find some indication that consumption of S. frutescens might be having a discernable impact on in vivo production of a key pro-inflammatory mediator, tumor necrosis factor-α (TNF-α). When otherwise healthy mice were fed a diet containing S. frutescens they showed a trend (i.e., p < 0.09) to produce nearly 50% less TNF-α following an injection of E. coli endotoxin (i.e., LPS). One might expect that any treatment that reduces in vivo production of this inflammatory mediator would be associated with notable changes in sickness behavior. [29] To examine this possibility we used voluntary locomotion as a surrogate for sickness behavior as described elsewhere. [29] Yet, we did not observed any indication that S. frutescens feeding affected voluntary locomotion prior to and during an active infection from either E. coli or during an infection from L. monocytogenes. We measured many cytokines and chemokines that play important roles in shaping the early immune response to infection, yet ex vivo production of IL-1α was the only one that was significantly affected by dietary S. frutescens consumption. Recently, Africa and Smith [30] reported that treatment of co-cultures of human monocytes and endothelial cells with an aqueous extract of S. frutescens greatly diminished the production of IL-1 in response to stimulation with HIV-1 antigens. That a recent human clinical trial found that consumption of milled vegetative parts of S. frutescens in capsules might promote tuberculosis, [31] is consistent with what would be expected if host production of TNF-α and IL-1β was diminished. [32,33] The present study was limited in a number of ways. First, the amount of S. frutescens included in the experimental diets may have been too low to elicit an observable effect. While we sought to use human equivalent dosing calculations derived from the research literature we may have been too conservative in setting our diet inclusion levels at 0.25 and 1.0%, by weight. It is possible that inclusion of more S. frutescens may have had a greater impact on in vivo host immune/inflammatory responses. Excessive production of inflammatory mediators such as ROS, NOS, and cytokines can result in tissue damage and even death. [34,35] Consistent with this was our preliminary findings that mice fed a diet containing 2% (wt/wt) of ground leaves from S. frutescens experienced lower survival rates (i.e., 17%, 1 of 6 mice survived) than mice fed a diet devoid of this botanical following an LD 50 E. coli challenge.
A second limitation emerged based on the contradictory findings generated in vitro with this botanical. We have reported that ethanolic extracts of S. frutescens possess potent antiinflammatory activities [36], while aqueous extracts of S. frutescens stimulate macrophage inflammatory responses. [1] Therefore, inclusion of unfractionated botanical in the diet may have failed to noticeably modify host response to bacterial challenge due to a cancelling effect of two or more components with opposite or contradictory activities. In an attempt to address this possibility, we conducted a small follow-up experiment where we administered by oral gavage either an aqueous or ethanolic extract of S. frutescens to a limited number of mice. After 7 days, half of the mice in each group were challenged with LPS or Pam2CSK and blood was collected 2 hours later. However, we found no discernible effect of either extract of S. frutescens on in vivo production of inflammatory cytokines and chemokines.
Third, our experimental design was limited in that we only included single time points for the assessment of bacterial clearance and circulating inflammatory cytokines and chemokines. It is possible that S. frutescens may have had a significant impact on host responses at different times after the immune challenges. Another obvious limitation of our study was that we only examined host response to a single gram-negative bacterium and single gram-positive bacterium (i.e., E. coli and L. monocytogenes, respectively). Host response to other pathogens, bacterial as well as viral, can vary tremendously. In short, our failure to observe significant modulation of host response to these two pathogens cannot exclude the possibility that treatment with S. frutescens might not provide benefits for host immunity against other infectious agents.

Conclusions
In conclusion, this study is the first to examine whether oral consumption of unfractionated vegetative parts of S. frutescens can modify the host immune response in vivo. We found that host clearance of a gram-positive and a gram-negative bacterium from mice was not significantly altered by S. frutescens intake. S. frutescens exhibited only a modest impact on the production of a couple of select inflammatory cytokines. Whether such changes occur in humans, and if they do, whether they would provide clinically relevant benefits to individuals using this medicinal plant remains uncertain.
Supporting Information S1 Fig. Impact of Dietary S. frutescens on Spontaneous Activity (i.e., sickness behavior) of Male Mice After an Experimental Infection with either L. monocytogenes. Healthy male BALB/c weanling mice were fed experimental diets containing one of three doses of S. frutescens (i.e., 0, 0.25 or 1% by wt) for 3-4 wks. Mice were housed in pairs in a vivarium with a 12 hr light:dark cycle at a room temperature between 22-25°C and a relative humidity of 50-60% for the entirety of the study. One of each pair of mice had a mini-emitter surgically implanted under the skin along their spine between their shoulders. At~1 week post-surgery mice received an injection of 10 4 cfu of L. monocytogenes, EGD strain. Each mouse's movement/ activity was recorded every 5 seconds and reported as accumulated events for every 20 minute period. Each tick mark on the X-axis represents an hour. Data shown represent the mean activity (#events) of four mice in each diet treatment group (n = 4/diet treatment group). (TIF) S2 Fig. Impact of Dietary S. frutescens on Spontaneous Activity (i.e., sickness behavior) of Male Mice After an Experimental Infection with E. coli. Healthy male C57BL/6 weanling mice were fed experimental diets containing one of three doses of S. frutescens (i.e., 0, 0.25 or 1% by wt) for 3-4 wks. Mice were housed in pairs in a vivarium with a 12 hr light:dark cycle at a room temperature between 22-25°C and a relative humidity of 50-60% for the entirety of the study. One of each pair of mice had a mini-emitter surgically implanted under the skin along their spine between their shoulders. At~1 week post-surgery mice received an injection of 10 8 cfu of E. coli K12 strain. Each mouse's movement/activity was recorded every 5 seconds and reported as accumulated events for every 20 minute period. Each tick mark on the X-axis represents an hour. Data shown represent the mean activity (#events) of four mice in each diet treatment group (n = 4/diet treatment group). (TIF) S1 Table. Impact of Dietary S. frutescens on Body, Liver, and Spleen Weight of Mice Following an L. monocytogenes challenge. Healthy female and male BALB/c weanling mice were fed experimental diets containing one of three doses of S. frutescens (i.e., 0, 0.25 or 1% by wt) for 3-4 wks. At~7 wk of age, mice were weighed (i.e., pre-challenge), then injected intravenously with~10 4 cfu of L. monocytogenes, EGD strain. Three days following the challenge, mice were re-weighed (i.e., post-challenge), then humanely killed for the collection of liver and spleen, which were weighed and then homogenized for the subsequent enumeration of bacteria. All values are expressed as means ± SEM (n = 15-16 per dietary treatment group). (DOCX) S2 Table. Impact of Dietary S. frutescens on Body, Liver, and Spleen Weight of Mice Following an E. coli challenge. Healthy female and male C57Bl/6 weanling mice were fed experimental diets containing one of three doses of S. frutescens (i.e., 0, 0.25 or 1% by wt) for 3-4 wks. At~7 wk of age, mice were weighed (i.e., pre-challenge), then injected intravenously with 10 8 cfu of E. coli, K12 strain. Two days following the challenge, mice were re-weighed (i.e., post-challenge), then humanely killed for the collection of liver and spleen, which were weighed and then homogenized for the subsequent enumeration of bacteria. All values are expressed as means ± SEM (n = 15-16 per dietary treatment group). (DOCX) S3 Table. Serum Cytokines and Chemokines from Mice 2 hours Following an Injection with Vehicle (PBS). For this study, male C57Bl/6 mice were fed experimental diets containing one of three doses of S. frutescens (0, 0.25 and 1% SF by wt) for 3 wks. Mice were injected at 1 min intervals with 1.0 mL of sterile PBS (i.e., vehicle controls for LPS challenged mice). Two hours following this injection blood was collected from mice and allowed to clot for 30 minutes at room temperature. Serum was collected by centrifugation and stored at -80°C until assayed for cytokines/chemokines using a commercial multiplex kit. Data shown are from eight mice from each diet treatment group (n = 8/trt); values represent means ± SEM (pg/mL). Diet intervention/treatment, as tested by ANOVA, failed to significantly affect any measured parameter (i.e., p > 0.05). (DOCX) S4 Table. Ex Vivo LPS-induced Inflammatory Cytokine and Chemokine Production by Primary Macrophages Isolated from Mice Fed Differing Levels of S. frutescens. For this experiment, male C57BL/6 mice were fed experimental diets containing one of three doses of S. frutescens (0, 0.25 and 1% by wt) for 3-4 wks. Peritoneal macrophages were isolated 3 days following intraperitoneal injection with sterile thioglycolate broth. Adherent cells (i.e., >95% macrophages) were co-cultured with 100 ng/mL of LPS (from E. coli 0111:B4) and 24 h cell later culture supernatants were collected and subsequently diluted 10-fold in FBS-free DMEM, then analyzed for specific cytokines and chemokines using a commercial multiplex beadsbased assay system. Data shown are from twelve mice from each diet treatment group (n = 11-12/trt); all values are LSmeans ± pooled SEM, expressed in pg/mL, unless otherwise indicated. (DOCX) S5 Table. Ex Vivo Pam2CSK-induced Inflammatory Cytokine and Chemokine Production by Primary Macrophages Isolated from Mice Fed Differing Levels of S. frutescens. For this experiment, male C57BL/6 mice were fed experimental diets containing one of three doses of S. frutescens (0, 0.25 and 1% by wt) for 3-4 wks. Peritoneal macrophages were isolated 3 days following intraperitoneal injection with sterile thioglycolate broth. Adherent cells (i.e., >95% macrophages) were co-cultured with 5 ng of Pam2CSK and 24 h later cell culture supernatants were collected and subsequently diluted 10-fold in FBS-free DMEM, then analyzed for specific cytokines and chemokines using a commercial multiplex beads-based assay system. Data shown are from twelve mice from each diet treatment group (n = 12/trt); all values are LSmeans ± pooled SEM, expressed in pg/mL, unless otherwise indicated.


text:
Membrane palmitoylated protein 2 is a synaptic scaffold protein required for synaptic SK2-containing channel function

Mouse CA1 pyramidal neurons express apamin-sensitive SK2-containing channels in the post-synaptic membrane, positioned close to NMDA-type (N-methyl-D-aspartate) glutamate receptors. Activated by synaptically evoked NMDAR-dependent Ca2+ influx, the synaptic SK2-containing channels modulate excitatory post-synaptic responses and the induction of synaptic plasticity. In addition, their activity- and protein kinase A-dependent trafficking contributes to expression of long-term potentiation (LTP). We have identified a novel synaptic scaffold, MPP2 (membrane palmitoylated protein 2; p55), a member of the membrane-associated guanylate kinase (MAGUK) family that interacts with SK2-containing channels. MPP2 and SK2 co-immunopurified from mouse brain, and co-immunoprecipitated when they were co-expressed in HEK293 cells. MPP2 is highly expressed in the post-synaptic density of dendritic spines on CA1 pyramidal neurons. Knocking down MPP2 expression selectively abolished the SK2-containing channel contribution to synaptic responses and decreased LTP. Thus, MPP2 is a novel synaptic scaffold that is required for proper synaptic localization and function of SK2-containing channels. DOI: http://dx.doi.org/10.7554/eLife.12637.001

Introduction
At most excitatory synapses in the central nervous system, such as the Schaffer collateral to CA1 synapses in the stratum radiatum of the hippocampus, excitatory neurotransmission is largely mediated by ionotropic AMPA-type (a-Amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid) and NMDA-type glutamate receptors. Yet, an emerging theme is that several conductances that limit membrane depolarization also make substantial contributions to the integrated excitatory post-synaptic potential (EPSP). For example, synaptically evoked Ca 2+ influx into dendritic spines activates apamin-sensitive SK2-containing channels (small conductance Ca 2+ -activated K + channels type 2; KCNN2), and their outward K + conductance shunts the AMPAR-mediated depolarization, effectively reducing the EPSP (Ngo-Anh et al., 2005;Faber et al., 2005). Kv4.2-containing channels are expressed in spines, close to, but not in the PSD (Kim et al., 2007). Synaptic activity evokes Ca 2+ influx through R-type voltage-gated Ca 2+ channels in spines that boosts nearby Kv4.2-containing A-type K + channels to further decrease the AMPA-mediated depolarization (Wang et al., 2014). In addition, Ca 2+ -activated Clchannels are expressed in the spines and provide further inhibitory contributions (Huang et al., 2012). Indeed, the sum of these repolarizing conductances may reduce the depolarizing AMPA-NMDA component by more than 50%. It is likely that each of these components can be regulated by a variety of second messenger pathways, greatly expanding the repertoire of targets to fine-tune synaptic transmission. For example, the Ca 2+ sensitivity of SK2 channels is regulated in an activity-dependent manner by co-assembled protein kinase CK2 and protein phosphatase 2A (Bildl et al., 2004;Allen et al., 2007) that are engaged by cholinergic signaling (Giessel and Sabatini, 2010). Moreover, the various contributions to synaptic responses may be dynamic, changing in response to distinct patterns of activity. Synaptic SK2-containing channels undergo protein kinase A (PKA)-dependent endocytosis upon the induction of LTP by theta burst pairing. The endocytosis of synaptic SK2-containing channels acts together with the PKA-dependent exocytosis of additional GluA1-containing AMPARs to mediate the expression of LTP . Moreover, after the initial expression of LTP and loss of the SK2-containing channel contribution, homeostatic mechanisms act to re-establish the synaptic SK balance (Lin et al., 2010). Similarly, Kv4.2-containing channels expressed in spines undergo PKA-dependent endocytosis after the induction of LTP (Kim et al., 2007;Hammond et al., 2008). Therefore, the appropriate localization, spatial distribution, and orchestrated dynamics of these protein complexes provide a powerful regulator of excitatory neurotransmission and plasticity.
One class of proteins that plays a major role in synaptic organization and dynamics are the MAGUKs (Elias and Nicoll, 2007), of which there are 10 subfamilies. These modular, usually multivalent scaffolds bind to synaptic receptors, channels, and signaling molecules to anchor them into their proper locations within the post-synaptic membrane (Oliva et al., 2012), creating a spatially and temporally restricted signaling domain (Hammond et al., 2008;Colledge et al., 2000;Dell'Acqua et al., 2006). Thus, within the post-synaptic density of excitatory synapses PSD-95 binds to NMDARs (Cousins and Stephenson, 2012), while SAP97 binds to AMPARs (Howard et al., 2010;Leonard et al., 1998), and Shank and Homer may serve as modular organizers of the lattice of synaptic MAGUKs (Sheng and Kim, 2000;Hayashi et al., 2009). However, the molecular mechanisms that engender synaptic localization and dynamics to SK2-containing channels are not well understood. There are two major isoforms of SK2 that are expressed in CA1 pyramidal neurons; SK2-L (long) has an extended intracellular N-terminal domain compared to SK2-S (short) and the two isoforms co-assemble into heteromeric channels (Strassmaier et al., 2005). In mice that selectively lack SK2-L expression, the SK2-S channels are expressed in the plasma membrane of dendrites and eLife digest The neurons in the brain communicate with each other by releasing chemical messengers across structures called synapses. This signaling always occurs in the same direction: at a given synapse, one neuron sends signals that bind to receptor proteins on the surface of the receiving neuron. Repeatedly signaling across a synapse strengthens it, making it easier to communicate across, and sometimes such stimulation can cause a persistent strengthening of the synapse: this is known as long-term potentiation. Changes in synaptic strength are important for learning and memory.
In the synapses formed between a type of brain cell called CA1 neurons, a protein called SK2 forms part of an ion channel in the membrane of the receiving neuron and is important for synaptic strengthening and long-term potentiation. To work correctly, the SK2 channels must be precisely positioned at the synapse, but the mechanisms responsible for this positioning were not clear. Now, by experimenting with purified proteins taken from the CA1 neurons of mice, Kim et al. show that SK2 physically interacts with a scaffold protein called MPP2. Further experiments revealed that MPP2 is responsible for positioning SK2 at the synapses, and this allows SK2-containing channels to contribute to long-term potentiation and synaptic strengthening.
During synaptic strengthening, it is possible that SK2 disengages from MPP2, which influences learning. The next step is to understand the processes that dictate this behavior. dendritic spines, yet fail to become incorporated into the post-synaptic membrane. Consequently, the SK2-containing channel contributions to EPSPs and plasticity are absent, and this loss of synaptic SK2-containing channel function enhances hippocampus-dependent learning tasks (Allen et al., 2011). To identify proteins that might serve to localize synaptic SK2-containing channels, candidate SK2 interacting proteins were identified. One of them, the MAGUK protein MPP2 (membrane palmitoylated protein 2), is localized to the PSD and is essential for synaptic SK2-containing channel function.

MPP2 interacts with SK2
Proteomic analyses (on high-resolution quantitative mass spectrometry) of SK2-containing channels immunoaffinity-purified from rodent whole brain membrane preparations suggested that the MAGUK protein, MPP2 might be an interaction partner. To further investigate this interaction, two newly generated antibodies targeting MPP2 were tested. Probing Western blots of proteins prepared from total mouse brain with either MPP2 antibody detected a predominant band at~55 kDa. Similarly a single band,~60 kDa, was detected in proteins prepared from HEK293 cells expressing FLAG-tagged MPP2, but not from mock transfected cells. For both brain and HEK293, pre-incubating the antibodies with the respective immunizing antigen abolished the bands ( Figure 1A,B).
Therefore, these two antibodies were used for affinity purifications combined with quantitative mass spectrometry (see Materials and methods). The results demonstrated that both antibodies robustly purified MPP2 and co-purified SK2 as well as SK3. In addition, these experiments identified DLG1 (SAP97) and Lin-homologs 7A and 7C as proteins co-assembling with MPP2 in rodent brain ( Figure 1C . To rule out off-target effects of the two MPP2 antibodies, we tested their binding of C8-tagged of SAP97, Lin7A, or Lin7C, C8-tagged versions of each of these proteins after expression in HEK293 cells ( Figure 1D). While all 8-tagged proteins were recognized by anti-C8 antibody only C8-MPP2 was recognized by anti-MPP2 antibodies ( Figure 1D). MPP2, a member of the p55 Stardust family of MAGUK proteins, is predicted to be a 552 amino acid protein. Like most MAGUKs, MPP2 is a modular protein comprised of several distinct protein-protein interaction domains. There are two predicted L27 domains, followed by a single predicted PDZ domain, and then the SH3-HOOK-GK domains.
To test for direct interaction with SK2 channels, SK2-L and SK2-S, the SK2 isoforms that contribute to synaptic SK2-containing channels (Allen et al., 2011), and either C8-MPP2 or PSD-95 were co-expressed in HEK293 cells. Immunoprecipitations were performed using an anti-SK2 antibody raised in guinea pig that is directed to the common C-terminal domain of the two SK2 isoforms, or using IgG as a control. Precipitated proteins were prepared as Western blots. Probing with anti-SK2 antibody that was raised in rabbits and directed against the same C-terminal sequence, demonstrated equivalent SK2 expression for input and after immunoprecipitation in each sample (Figure 2-figure supplement 1). Probing with anti-C8 antibody detected a band of the appropriate apparent molecular weight for C8-MPP2 in the sample co-expressed with SK2-S plus SK2-L but not in the IgG control sample (Figure 2A). Probing with anti-PSD-95 antibody did not detect co-immunoprecipitation with SK2 ( Figure 2B). The results demonstrated that C8-MPP2 but not PSD-95 was specifically co-precipitated with SK2. To examine the possibility that SAP97 interacts with SK2-S, C8-SAP97 was co-expressed in HEK293 cells together with either myc-SK2-S or GluA1, a known SAP97 interaction partner (Leonard et al., 1998). Immunoprecipitations were performed using anti-SK2 antibody, anti-GluA1 antibody, or IgG as a control. Western blotting with anti-C8 antibody revealed that C8-SAP97 only co-immunoprecipitated when co-expressed with GluA1, despite equivalent levels of input and immunoprecipitated proteins ( Figure 2-figure supplement 1). To test if MPP2 interacts with the unique N-terminal domain of SK2-L that is required for synaptic localization GST pull-down experiments were performed. While PDZ interactions are important for many MAGUKpartner protein interactions, the unique N-terminal domain of SK2-L does not contain a PDZ ligand motif and using the PDZ domain from MPP2 failed to show an interaction with the N-terminal domain of SK2-L. Therefore, the SH3-HOOK-GK domain of MPP2 was employed. This domain of MPP2 as well the SH3-HOOK-GK domains from SAP97, a MAGUK scaffold of a distinct subfamily Western blots were prepared using mouse brain homogenate (Brain; 100 mg) or HEK293 cell extracts (1% of 10 -cm plate), either transfected to express FLAG-MPP2 (HEK) or empty plasmid (HEK mock). Duplicate lanes were prepared and one set was probed with the indicated MPP2 antibody (left panels) while the second set was probed with the same antibody after pre-absorbing with the immunizing antigen (right panels). Native MPP2 and transfected FLAG-MPP2 were detected only in the left panels. (C) Bar graphs illustrating abundance ratios determined for the indicated proteins in APs with two anti-MPP2 antibodies and IgG as a negative control. Horizontal lines denote threshold for specificity of co-purification. (D) Western blots of Figure 1 continued on next page from MPP2 (Oliva et al., 2012), or the SH3-HOOK-GK domain from CaCNB4, a non-canonical MAGUK protein that is a beta subunit for voltage-gated Ca 2+ channels (Van Petegem et al., 2004) were prepared as GST-fusion proteins, bound to glutathione-agarose beads, and used as baits for the prey, the His-tagged N-terminal domain of SK2-L. A GST-fusion protein of the C-terminal domain of Kv1.4, and a His-fusion protein of PSD-93 (Chapsyn) were also prepared as positive interaction controls (Lunn et al., 2007). Coomassie staining and Western blotting with anti-GST antibody verified equivalent amounts of the baits, either as input or bound to beads (Figure 2-figure supplement 2). After exposure to the baits, bound prey proteins were eluted and prepared as Western blots. Probing with an anti-His antibody showed that the SH3-HOOK-GK domain from MPP2, but not from SAP97 or CaCNB4, specifically pulled down the N-terminal domain of SK2-L. As expected,  Figure 2C). Taken together these results suggest that MPP2 interacts with the unique N-terminal domain of SK2-L that is required for synaptic localization (Allen et al., 2011).

MPP2 is a synaptic MAGUK protein
To examine the anatomical localization of MPP2, the anti-MPP2 antibodies were used for immunohistochemistry on hippocampal sections. At the light level, MPP2 was expressed throughout the hippocampus, and in area CA1, MPP2 was predominantly expressed in the dendritic arbors ( Figure 3A, B). To determine the subcellular profile of MPP2 expression, pre-and post-embedding immunogold electron microscopy (iEM) was performed. Using pre-embedding techniques, sections from three animals revealed immunoparticles for MPP2 prominently labeled dendritic spines and dendritic shafts both along the plasma membrane and at intracellular sites. Post-synaptically, most immunoparticles for MPP2 were found in spines (84%; 436 particles) versus in dendrites (16%; 82 immunoparticles), with most immunoparticles in either compartment localized to the plasma membrane (84%; 365 immunoparticles in spines and 65%; 54 immunoparticles in dendrites). Next, post-embedding techniques were applied to determine if MPP2 is expressed in the post-synaptic density. Sections from three animals revealed that immunoparticles for MPP2 prominently labeled the postsynaptic densities of dendritic spines, as well as being detected in dendritic shafts ( Figure 3C,D). Immunoparticles for MPP2 were not detected in stratum pyramidale, and were also absent pre-synaptically. Therefore, MPP2 is a synaptic MAGUK protein.

MPP2 is required for synaptic SK2-containing channel function
To test whether MPP2 expression is important for synaptic SK2 channel function, two short hairpin RNAs (shRNAs) targeting the 3' untranslated region (3' UTR) of Mpp2 mRNA were co-expressed in area CA1 by in utero electroporation (e14-16) of a plasmid that also directed expression of the fluorescent protein, GFP. Four-to five-week-old mice were then used to prepare fresh hippocampal slices. Whole-cell current clamp recordings were made from CA1 pyramidal neurons, either transfected, as reported by GFP expression, or non-transfected control cells. Synaptic stimulations of the Schaffer collateral axons evoked EPSPs. After establishing a stable baseline, apamin (100 nM) was applied to the slices. For control cells, apamin increased EPSPs to 144.4 ± 4.3% of baseline (n = 15; p<0.0001), consistent with previous studies (Ngo-Anh et al., 2005;Lin et al., 2008;Giessel and Sabatini, 2011). In contrast, apamin did not significantly affect EPSPs from transfected CA1 pyramidal neurons (94.6 ± 3.3%; n = 14) ( Figure 4A-C).
To further verify the efficacy of MPP2 knock-down, pre-embedding iEM was performed on hippocampal sections from MPP2 shRNA-transfected animals. To identify transfected CA1 pyramidal neurons, mouse anti-GFP antibody was detected using HRP-linked anti-mouse secondary antibody, and rabbit anti-MPP2 antibody was detected using immunogold particles coupled to anti-rabbit secondary antibody. Immunoparticles were then counted in the same number of profiles belonging both to dendritic spines and dendritic shafts on single sections. In spines on neurons expressing GFP, immunoparticles for MPP2 were reduced by 81% compared to spines on non-transfected control neurons (GFP-negative: 417 particles in 60 spines; GFP-positive: 101 particles in 60 spines; p<0.001). A similar reduction (78%) was seen in dendritic shafts (GFP-negative: 155 particles in 25 dendrites; GFPpositive: 44 particles in 25 dendrites; p<0.001) ( Figure 5A-D).
To demonstrate that the effects of shRNA transfection were specifically due to MPP2 knockdown, the shRNA (GFP) plasmid was co-transfected with a plasmid that directed expression of shRNA-immune MPP2 and a plasmid that expressed the red fluorophore, mApple. In doubly transfected neurons, apamin increased EPSPs similar to control, non-transfected neurons (148.2 ± 4.2%; n = 13; p<0.0001, compared to shRNA knock-down) ( Figure 6A,B). Thus, the consequences of the shRNA expression are mediated by knock-down of MPP2.

MPP2 selectively affects synaptic SK2-containing channels
There are at least two functionally distinct populations of apamin-sensitive SK2-containing channels in CA1 pyramidal neurons. One population resides in the spines and is activated by synaptically evoked NMDAR-dependent Ca 2+ influx (Ngo-Anh et al., 2005). Another population resides in the dendrites and may be activated by somatic voltage steps that induce Ca 2+ influx through voltagegated Ca 2+ channels (Stocker et al., 1999;Gerlach et al., 2004;Bond et al., 2004). To determine if MPP2 knock-down specifically affects synaptic SK2-containing channels or also affects SK2-containing channels in the dendrites transfected or control neurons were clamped at À55 mV and stepped to 20 mV; tail currents were elicited upon stepping back to À55 mV ( Figure 6A). The apamin-sensitive component of the tail current ( Figure 7A inset) effectively measures the SK2-containing channel component activated by somatic depolarization and showed that MPP2 knock-down did not significantly affect somatic activation of SK2-containing channels (ctrl: 41.9 ± 8.3 pA, n = 13; transfected: 53.7 ± 10.5, n = 7; P = 0.44) ( Figure 7A,B). These results show that knocking down MPP2 expression Figure 5. Efficient knock-down of MPP2 expression in CA1 pyramidal neurons. Immunoreactivity for MPP2 in the CA1 region of the hippocampus, as revealed using a double-labelling pre-embedding method. (A-D) The peroxidase reaction end product (HRP) indicating GFP immunoreactivity filled CA1 pyramidal cells, whereas immunoparticles for MPP2 were mainly located along the plasma membrane and at intracellular sites of pyramidal cells. Immunoparticles for MPP2 were distributed in both GFP-positive (crossed arrows) and GFP-negative (arrows) dendritic spines (s) and dendritic shafts (Den) of pyramidal cells. However, there was a striking reduction of immunoparticles for MPP2 in GFP-positive profiles compared to GFP-negative profiles (see text). at, axon terminal. Scale bars in A-D: 0.2 mm. DOI: 10.7554/eLife.12637.011 disrupts the synaptic SK2-containing channel component but does not affect the channels in the dendrites that are activated by somatic voltage pulses.

Loss of MPP2 reduces LTP
SK channel activity modulates the induction of synaptic plasticity (Stackman et al., 2002) and the activity-dependent endocytosis of synaptic SK2-containing channels contributes to the expression of LTP . To determine the consequences of MPP2 knock-down and loss of synaptic SK2-containing channel activity on LTP, a theta burst pairing protocol was delivered in which Schaffer collateral stimulations were paired with back-propagating action potentials evoked by somatic current injections. In non-transfected CA1 pyramidal neurons, this induced LTP of 407.9 ± 46.3% (n = 8), while in transfected, MPP2 knock down neurons, LTP was significantly reduced (243.0 ± 14.8%; n = 13; p<0.001) (Figure 8). Consistent with the loss of synaptic SK2-containing channels, MPP2 knock-down reduces LTP.

Discussion
The results presented here identify the synaptic MAGUK protein, MPP2 (p55), that is required for synaptic SK2-containing channel function. Synaptically evoked EPSPs in CA1 pyramidal neurons are increased by the SK channel blocker, apamin, but in CA1 pyramidal neurons expressing shRNAs directed against Mpp2 mRNA apamin has no effect. MPP2 knockdown selectively affects synaptic SK2-containing channel function, as the SK2-containing channels expressed in the dendrites that are activated by somatic voltage steps are not altered. Consistent with the effects of MPP2 on synaptic SK2-containing channels, MPP2 knock-down reduces the expression of TBP-induced LTP by~30%. This is slightly more than the component of LTP attributed to SK2 endocytosis in untransfected CA1 pyramidal neurons,~17% . This might reflect effects of MPP2 knock-down on other interaction partners (see below).
Previous results showed that synaptic SK2-containing channels are heteromeric assemblies that contain two isoforms of SK2, SK2-S and SK2-L. Compared to SK2-S, SK2-L has an additional 207 amino acids in the intracellular N-terminal domain and SK2-S is otherwise entirely contained in SK2-L (Strassmaier et al., 2005). In a transgenic mouse selectively lacking SK2-L, the SK2-S channels are expressed in the dendrites and even in the plasma membrane of dendritic spines, but they are specifically excluded from the PSD, and apamin fails to boost synaptically evoked EPSPs. Re-expressing SK2-L reinstates synaptic function as measured by apamin sensitivity of EPSPs (Allen et al., 2011). These results implicated the unique N-terminal domain of SK2-L in directing SK2-containing channel synaptic localization and function, and suggested that the N-terminal domain of SK2-L might interact with a partner protein to engender PSD localization. Indeed, MPP2 binds to the unique N-terminal domain and knocking down MPP2 expression phenocopied the effect of SK2-L deletion on synaptic responses. MAGUK proteins recruit and stabilize AMPA and NMDA receptors in the PSD. Our results suggest that MPP2 similarly serves to stabilize SK2-containing channels in the PSD.
MPP2 is a member of the p55 Stardust subfamily of MAGUK scaffold proteins, named after the founding member MPP1, the major palmitoylated protein in erythrocytes (Alloisio et al., 1993). Similar to other MAGUK scaffold proteins, MPP2 is modular, consisting of two L27 domains that may mediate homo-or heterophilic interactions, a single PDZ domain followed by SH3-HOOK-GK domains. Biochemical studies showed that MPP2 is enriched in the PSD fractions of rat brain, and pull-down assays to test for interactions suggested MPP2 may interact with itself as well as a number of other synaptic proteins, among them are PSD-95, SAP97, GKAP, CASK, GRIP, neuroligin, and CaMKII. The SH3-HOOK-GK domain of MPP2 was implicated in mediating these interactions (Jing-Ping et al., 2005), similar to the interaction between MPP2 and the N-terminal domain of SK2-L.
The results presented here, using an unbiased approach also identified SK3 as immunopurifying with MPP2. SK2 and SK3 can form heteromeric channels in brain (Strassmaier et al., 2005), and SK3 is expressed in CA1 pyramidal neurons (Ballesteros-Merino et al., 2014). Different from SK2 that has two N-terminal isoforms, there is only one SK3 N-terminal isoform and it is similar to the extended N-terminal domain of SK2-L, harboring several islands of homology that might mediate interactions with MPP2.
The proteomics analyses of MPP2 also identified three additional scaffold proteins, the MAGUK protein, DLG1 (SAP97), as well as Lin7A and Lin7C as MPP2-interacting proteins. In epithelial cells, MPP7, a closely related member of the p55 Stardust family, dimerizes with Lin7 proteins, an interaction mediated by the C-terminal L27 domain, and the dimeric complex then associates with DLG1 via the N-terminal L27 domain that is insufficient to mediate DLG1 interactions in the absence of bound Lin7 (Bohl et al., 2007). DLG1 also binds the C-terminal PDZ ligand on the GluA1 subunit of AMPA receptors (Leonard et al., 1998). Indeed, upon the induction of LTP at Schaffer collateral to CA1 synapses, additional GluA1-containing AMPA receptors undergo exocytosis at a perisynaptic site followed by translocation into the post-synaptic membrane, increasing the AMPA component of EPSPs (Yang et al., 2008). This exocytosis is dependent on the PDZ ligand at the C-terminus of the GluA1 subunit (Lin et al., 2010;Yang et al., 2008;Shi et al., 2001), and exocytosis of GluA1-containing AMPA receptors is prerequisite to SK2-containing channel endocytosis; specifically blocking GluA1-containing AMPA receptor exocytosis prevents the rapid, subsequent endocytosis of SK2containing channels (Lin et al., 2010). Moreover, immunopurification of AMPA receptors from whole brain identified MPP2 as a protein that co-purified with AMPA receptors (Schwenk et al., 2012). Synaptic SK2-containing channels reside in very close proximity to synaptic NMDA receptors within the PSD, providing a molecular microdomain that facilitates their functional coupling (Ngo-Anh et al., 2005;Lin et al., 2008). PSD-95 interacts with NMDARs (Kornau et al., 1995), is crucial for the proper synaptic localization of ionotropic glutamate receptors (Schnell et al., 2002;Elias et al., 2008), and interacts with SAP97 (Cai et al., 2006). It will be interesting to determine whether MPP2 additionally interacts, directly or indirectly with PSD-95 to maintain the spatial synaptic relationship between SK2-containing channels and NMDA receptors. MPP2 contains multiple different protein-protein interaction domains that may bind not only receptors and channels but, additionally, signaling molecules as well as connections to the cytoskeleton. These observations raise the possibility that there is a dynamic protein lattice encompassing SK2-containing channels, AMPARs and NMDARs, and regulatory proteins that is woven together by molecular interactions between scaffold proteins to precisely tune synaptic responses during basal neurotransmission and plasticity.

Animal care
All procedures involving animals were performed in accordance with the guidelines of Oregon Health and Science University (Portland, OR), animal care protocol number: IP00000191; University of Freiburg (Freiburg, Germany), Regierungsprä sidium Freiburg, AZ: 35-9185/G-12/47; and University of Castilla-La Mancha (Albacete, Spain).

Molecular biology
MPP2 shRNAs were designed using an online algorithm (http://sirna.wi.mit.edu). Two sequences targeting the 3'UTR of mouse Mpp2 mRNA (NM_016695; 1811-1833; 3620-3642) were synthesized for shRNA expression and cloned into a vector that drove their expression from the U6 promoter. This same plasmid also directed GFP expression from the ubiquitin promoter. For rescuing MPP2 knockdown, the MPP2 shRNA plasmids (GFP) were co-transfected with a plasmid expressing the MPP2 coding sequence, cloned between the CAG promoter and the 3' UTR from bovine growth hormone. This plasmid was co-transfected with a plasmid directing mApple expression from the ubiquitin promoter.

Antibodies
To generate antibodies to MPP2, rabbits were immunized with synthetic peptides representing amino acids 116-145 and 480-507 of mouse MPP2 (NP_057904). To test the antibodies using Western blots, mouse whole brains were homogenized in ice-cold homogenization buffer (0.32 M sucrose, 1 mM ethylenediaminetetraacetic acid, 1 mM ethyleneglycoltetraacetic acid, 10 mM Tris-HCl, pH 7.2, 0.4 mM phenylmethylsulfonyl fluoride). The resultant homogenate was subjected to centrifugation at 1000 g for 10 min to remove nuclei and debris. The protein concentration was determined with Lowry's method. Human embryonic kidney 293T (HEK293T) cells were transfected with pFLAG-CMV vector (Clontech, Palo Alto, CA) encoding mouse FLAG-MPP2 in a 10 cm dish. Cells were harvested in 0.5 ml of PBS. Homogenates and suspended cells were mixed with an equal volume of 2 x sodium dodecyl sulfate (SDS) sampling buffer (63 mM Tris-HCl, pH 6.8, 4% SDS, 20% glycerol, 0.002% bromophenol blue), and denatured with 50 mM ( ± )-dithiothreitol at 55˚C for 30 min. Proteins (100 mg of brain homogenates and 0.5 ml of cell lysates) were separated using 10% SDS-polyacrylamide gel electrophoresis, and electroblotted onto an Immobilon-P Transfer Membrane (Millipore, Billerica, MA). After blocking with 5% skimmed milk for 30 min, blotted membranes were incubated with primary antibodies (1 mg/ml) for 1 hr, then with peroxidase-conjugated secondary antibodies for 1 hr (Jackson ImmunoResearch, West Grove, PA; 1:10,000). Tris-buffered saline (10 mM Tris-HCl, pH 7.5, 150 mM NaCl) containing 0.1% Tween-20 was used as the dilution and washing buffer. Immunoreactions were visualized with the ECL chemiluminescence detection system, and captured using an ImageQuant LAS 500 (GE Healthcare, Buckinghamshire, UK). For specificity control, anti-MPP2a and anti-MPP2b (1 mg/ml) were mixed with 20 mg/ml of GST fusion proteins of the respective immunizing antigen.

Proteomic analysis
Proteomic analysis of MPP2 from rat brain was performed as described previously (Schwenk et al., 2012;Schwenk et al., 2010;Bildl et al., 2012). Briefly, affinity-purifications (APs) with anti-MPP2a, anti-MPP2b, and pre-immunization immunoglobulins (IgG) were performed on rat brain membrane fractions solubilized with CL-91 (Logopharm, March, Germany) and whole eluates were subjected to high-resolution mass spectrometry. Proteins retained in APs and identified by mass spectrometry were quantified by integration of peptide m/z signal intensities over time (peak volumes, PVs) that were extracted from FT full scans using MaxQuant (v.1.4.1.2; http://www.maxquant.org; with integrated off-line mass calibration). Relative abundance of proteins in anti-MPP2 samples versus control (abundance ratio or rPV) was determined by the TopCorr method  as the median of either (i) six individual peptide PV ratios of the best correlating protein-specific peptides (as determined by Pearson correlation of their abundance values; TopCorr6), or (ii) the 50% best of all individual peptide PV ratios (Top50). The linear dynamic range of the TopCorr method is about 4 orders of magnitude (detailed in ). The data used for rPV determination (including peptide sequences, PV and rPV values, as well as the respective medians and selections methods) are summarized in Supplementary file 1. The coverage of the primary sequences of all proteins shown in Figure 1C is presented in Figure 1

Co-immunoprecipitation
HEK293 cells were transfected using Lipofectamine 2000 (Thermo Scientific). After 48 hr cells were washed, collected with PBS buffer, and pelleted. Cells were then solubilized with lysis buffer (20 mM HEPES, pH 7.5, 150 mM NaCl, 10% glycerol, 2 mM EDTA, 1 mM PMSF, and protease inhibitors (Hoffman La Roche, Basel, Switzerland) containing 1% b-D-dodecyl maltoside (Sigma-Aldrich, St. Louis, MO) for 30 min at RT. The lysate was centrifuged at 14,000 rpm for 20 min at 4˚C. The supernatant was incubated with the indicated primary antibody for 30 min then 20 ml of protein A/G agarose beads (Thermo Scientific) were added for incubation overnight at 4˚C with rotation. The beads were washed three times in washing buffer (mM) (20 HEPES, pH 7.5, 150 NaCl, 10 KCl, 2 EDTA, 10% glycerol). Proteins were eluted with 40 ml of 2X SDS sample buffer at 37˚C. Bound and eluted proteins were subsequently separated by SDS-PAGE and transferred to PVDF membrane (BioRad, Hercules, CA). After blocking with 5% skimmed milk for 1 hr, the membrane was probed with the indicated antibodies over night at 4˚C. HRP-conjugated secondary antibodies were applied for 30 min at RT. Blots were detected with SuperSignal ECL (Thermo Scientific) and developed with GeneMate Blue film (BioExpress, Kaysville, UT).

Immunoelectron microscopy
Immunohistochemical reactions at the electron microscopic level were carried out using immunogold methods as described previously (Luján et al., 1996).

Pre-embedding immunohistochemistry
Single-and double-labelling methods were used. Briefly, in single-labelling experiments, free-floating sections were incubated in 10% NGS diluted in TBS. Sections were then incubated in anti-MPP2 antibodies (1-2 mg/ml diluted in TBS containing 1% NGS), followed by incubation in goat anti-rabbit IgG coupled to 1.4 nm gold (Nanoprobes, Yaphank, NY). Sections were postfixed in 1% glutaraldehyde and washed in double-distilled water, followed by silver enhancement of the gold particles with an HQ Silver kit (Nanoprobes). In co-labelling experiments, GFP immunoreactivity was visualized by the immunoperoxidase reaction, and MPP2 immunoreactivity was revealed with the silver-intensified immunogold reaction. Sections were then treated with osmium tetraoxide (1% in 0.1 M PB), block-stained with uranyl acetate, dehydrated in graded series of ethanol and flat-embedded on glass slides in Durcupan (Sigma-Aldrich) resin. Regions of interest were cut at 70-90 nm on an ultramicrotome (Reichert Ultracut E, Leica Biosystems, Barcelona, Spain) and collected on 200-mesh copper grids. Staining was performed on drops of 1% aqueous uranyl acetate followed by Reynolds's lead citrate.

Post-embedding immunohistochemistry
Briefly, ultrathin 80-nm-thick sections from Lowicryl-embedded blocks of the hippocampus were picked up on coated nickel grids and incubated on drops of a blocking solution consisting of 2% human serum albumin in 0.05 M TBS and 0.03% Triton X-100. The grids were incubated with MPP2 antibodies (10 mg/ml in 0.05 M TBS and 0.03% Triton X-100 with 2% human serum albumin) at 28˚C overnight. The grids were incubated on drops of goat anti-rabbit IgG conjugated to 10 nm colloidal gold particles (Nanoprobes) in 2% human serum albumin and 0.5% polyethylene glycol in 0.05 M TBS and 0.03% Triton X-100. The grids were then washed in TBS and counterstained for electron microscopy with saturated aqueous uranyl acetate followed by lead citrate.
Ultrastructural analyses were performed in a Jeol-1010 electron microscope (Synaptic Structure Laboratory, School of Medicine, University of Castilla-La Mancha, Albacete, Spain). Electron photomicrographs were captured with an ORIUS SC600B CCD camera (Gatan, Munich, Germany). Digitized electron images were modified for color, brightness, and contrast with Adobe Photoshop, version 7.0. Labelled structures were classified based on unambiguous morphological information in each section. Axon terminals were identified by the presence of synapses and small round and/or large granular vesicles. Synapses were identified as parallel membranes separated by widened clefts that were associated with membrane specializations. Synapses displaying a prominent density on the post-synaptic side were characterized as asymmetrical. Dendritic spines were identified as small protrusions exhibiting membrane continuity with the dendritic shaft. Significance of immunogold particle labelling in GFP-positive versus GFP-negative spines and dendrites was assessed by ANOVA.

In utero electroporation
Timed-pregnant mice were anesthetized with isofluorane, their abdominal cavity cut open, and the uterine horns/sac exposed. Approximately 2 ml of DNA solution (~2 mg/ml) was injected into the lateral ventricle of e14-e16 embryos, using a glass pipet pulled from thin walled capillary glass (TW150F-4, World Precision Instruments, Sarasota, FL) and a Picospritzer III microinjection system (Parker Hannifin, Hollix, NH). The head of each embryo within its uterine sac was positioned between tweezer-type electrodes (CUY650P10; Sonidel, Dublin, Ireland), and 5 square electric pulses (35 V; 50 ms; 1-s intervals) were passed using an electroporator (CUY21; Sonidel). After electroporation, the wall and skin of the abdominal cavity of the pregnant mouse was sutured-closed, and embryos were allowed to develop normally.
Electrophysiology CA1 pyramidal cells were visualized with IR/DIC optics (Olympus BX51W1; Olympus Scientific Solutions, Waltham, MA) and a CCD camera. Whole-cell, patch-clamp recordings were obtained from CA1 pyramidal cells using a Multiclamp 700B (Molecular Devices, Sunnyvale, CA), digitized using an ITC-18 analog-to-digital converter, and transferred to a computer using Patchmaster software (Heka Instruments, Bellmore, NY). Patch pipettes (open pipette resistance, 2-4 MW) were filled with (in mM) 133 K-gluconate, 4 KCl, 4 NaCl, 1 MgCl 2 , 10 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), 4 MgATP, 0.3 Na 3 guanosine triphosphate (Na 3 GTP), and 10 K 2 -phosphocreatine (pH 7.3). Electrophysiological records were filtered at 3.3 kHz and sampled at 10 kHz. Series resistance was electronically compensated to greater than 70%. A bias current was applied to maintain the membrane potential in current clamp at À65 mV. The input and residual series resistance in current clamp was determined from a 20 pA hyperpolarizing pulse applied at the end of each sweep. The input resistance in voltage clamp was determined from a 5 mV hyperpolarizing pulse applied at the beginning of each sweep. All recordings were from cells with a resting membrane potential less than À60 mV and a stable input resistance. All electrophysiological recordings were performed at 22-24˚C, and data were not corrected for a junction potential of~15 mV of the internal solution with respect to the bath ACSF.

Synaptic stimulation
EPSPs were recorded in whole-cell mode. Capillary glass pipettes (tip diameter,~5 mm) filled with ACSF and connected to Digitimer constant current stimulus isolation unit (AutoMate Scientific, Berkeley, CA) were used to stimulate pre-synaptic axons in stratum radiatum as described in Results. SR95531 (2-10 mM) and CGP55845 (1 mM) were present to reduce GABA A and GABA B contributions, respectively. To prevent epileptic discharges in the presence of GABAergic blockers, the CA3 region was microdissected out of slices used for EPSP recordings. LTP was induced by a theta burst pairing protocol, as previously described .

Data analysis
Data were analyzed using IGOR (WaveMetrics, Lake Oswego, OR). Data are expressed as mean ± SEM. Paired two-sample t-tests were used to determine significance of data in the same pathway, and non-parametric Wilcoxon Mann-Whitney two-sample rank test was used to determine significance between groups of data; p<0.05 was considered significant.

Chemicals and solutions
D-AP5, CNQX, CGP55845, and SR95531 were obtained from Tocris Bioscience (Ellisville, MO). All other chemicals were obtained from Sigma-Aldrich unless specified. All perfusing solutions were modified from regular ACSF unless otherwise noted.