Emerging evidence indicates that metabolic programs regulate B cell activation and Ab responses. However, the metabolic mediators that support the durability of the memory B cell and long-lived plasma cell populations are not fully elucidated. Adenosine monophosphate–activated protein kinase (AMPK) is an evolutionary conserved serine/threonine kinase that integrates cellular energy status and nutrient availability to intracellular signaling and metabolic pathways. In this study, we use genetic mouse models to show that loss of ΑMPKα1 in B cells led to a weakened recall Ab response associated with a decline in the population of memory-phenotype B cells. AMPKα1-deficient memory B lymphocytes exhibited aberrant mitochondrial activity, decreased mitophagy, and increased lipid peroxidation. Moreover, loss of AMPKα1 in B lymphoblasts was associated with decreased mitochondrial spare respiratory capacity. Of note, AMPKα1 in B cells was dispensable for stability of the bone marrow–resident, long-lived plasma cell population, yet absence of this kinase led to increased rates of Ig production and elevated serum Ab concentrations elicited by primary immunization. Collectively, our findings fit a model in which AMPKα1 in B cells supports recall function of the memory B cell compartment by promoting mitochondrial homeostasis and longevity but restrains rates of Ig production.

B cells confer long-lasting protection against microbes or their exotoxins via the production of highly specific Abs. Upon encounter with cognate Ag, naive B cells rapidly proliferate and ultimately differentiate into Ag-specific memory B cells or Ab-secreting plasma cells, a subset of that persists for decades after clearance of the Ag (1). The persistence of memory B cell and long-lived plasma cell populations is critical for protection against subsequent Ag exposure and achieving long-lasting immunological memory.

After activated B cells undergo proliferation and differentiation, long-lived plasma cells and memory B cells arise from both germinal center (GC)–dependent and –independent pathways (2). After a phase as plasmablasts, long-lived plasma cells can reside in the bone marrow and continuously synthesize and release Ag-specific Ab into the blood stream to bind Ag (3). By contrast, Ag-specific memory B cells continue to circulate through the body and have the capacity to reenter GC or rapidly differentiate into Ab-secreting cells (ASCs) upon subsequent exposure to Ag (4). Although both are long-lived cells in the B lineage, memory B cells appear to be metabolically quiescent, whereas plasma cells continuously synthesize large amounts of secretory glycosylated Ab (5). Apart from diversity in function, long-lived B cells must also adapt to varied nutrient availability in the microenvironment (6, 7). Mechanisms that support longevity in the face of metabolic stressors and demands are incompletely understood.

Bone marrow plasma cell longevity is attributable in part to the expression of Bcl6-related transcription factor ZBTB20 and the ability to shuttle pyruvate to the mitochondria (810). Emerging evidence implicates autophagy as critical for the long-term survival of both memory and plasma cells (1113). However, the mode of autophagy that supports the longevity of the B lineage is unknown.

The energy sensor adenosine monophosphate–activated protein kinase (AMPK) is one upstream activator of canonical autophagy. AMPK is a conserved serine/threonine kinase that couples nutrient availability to cellular metabolism (14). During periods of bioenergetic insufficiency, AMPK can restore cellular energy homeostasis by targeting multiple downstream targets that trigger pathways that increase ATP generation while inhibiting those that consume ATP. Specifically, AMPK promotes fatty acid oxidation, autophagy, and mitochondrial biogenesis while inhibiting protein and fatty acid synthesis (14). Several studies link age-related diseases to the dysregulation of mitochondrial dynamics, suggesting that AMPK may support longevity of cells through regulation of mitochondrial homeostasis (15). Furthermore, AMPK negatively regulates another energy sensor, mechanistic target of rapamycin complex 1 (mTORC1), which has been reported to be critical for effective primary Ab responses and the generation of memory (16).

Several reports indicate that AMPK activity in T cells is critical for effective activation and survival (17). In one study, AMPK supported the survival of the CD8 memory population; others showed defects in primary CD8 T cell activation and effector functions with the loss of AMPK (1820). In contrast to the T lineage, two studies have reported negative findings about AMPK in humoral responses (21, 22). B and T cells from mice with unconditional null alleles encoding the isoform AMPKα1 were sensitized to cell death upon metabolic stressors, but no effect on primary or boosted Ab responses was observed (21). However, later work showed that this whole-body knockout leads to diffuse inflammation as a consequence of AMPKα1 deficiency in other cell populations, including erythrocytes and APC, which may affect B cell function (23, 24). More recently, work with a B cell–intrinsic knockout of AMPKα1 concluded that loss of AMPK downregulates transcriptional expression of IgD (22). However, whether AMPK governs humoral memory remains unresolved.

Because AMPK is a master regulator that controls multiple aspects of metabolism, including mitochondrial homeostasis, mTORC1, and autophagy, all of which play critical roles in B cells fate and function, we sought to test if these downstream pathways were AMPK dependent in the B lineage. In this study, we show that AMPK promoted mitochondrial homeostasis and limited mTORC1 activity in this population while balancing effector functions and long-term maintenance of memory-phenotype B cells (MphenBC) as well as recall capacity after rechallenge. These findings provide evidence of a divergence between the cellular sources of two limbs in humoral memory—plasma cells and B cell memory—in their dependence on AMPK.

B cell–specific, AMPKα1-deficient mice were generated by crossing Prkaa1 flox mice, purchased from The Jackson Laboratory (Bar Harbor, ME), with mice harboring Cre recombinase under three types of control element: mb1-Cre (25), human CD20 (huCD20)–CreERT2, and Rosa26-CreERT2 (16). Age-matched Prkaa1+/+ mice with the corresponding Cre transgene were used as wild-type controls and cohoused with Prkaa1f/f mice. Mice were housed at Vanderbilt University under specified pathogen-free conditions, and both males and females (6–10 wk old) were used according to approved mouse Institutional Animal Care and Use Committee protocols. Because of practical limitations on mobilization of matched mice for immunization experiments with otherwise unmanipulated mice, the Rosa26-CreERT2 knock-in mice were used for purification of B cells for in vitro or transfer experiments as described below. For immunizations in mice with B lineage–specific loss of AMPKα1, huCD20-CreERT2 and mb1-Cre lines were immunized with 4-hydroxy-3-nitrophenylacetyl hapten conjugated to keyhole limpet hemocyanin (NP-KLH; Biosearch Technologies, Novato, CA) emulsified in Imject Alum (50 μg in 50 μl) (Thermo Fisher Scientific, Waltham, MA). Recall responses were induced with 100 μg NP-KLH in alum (100 μl). To induce deletion of Prkaa1 in mature B cells prior to immunization, mice with huCD20-CreERT2 or Rosa26-CreERT2 transgenes received three doses (3 mg each) of tamoxifen from Cayman Chemical (Ann Arbor, MI) dissolved in safflower oil injected i.p. every other day (16).

For transfer experiments, CD45.1+ IgHa recipient mice were generated by crossing CD45.1+ IgHb and CD45.2+ IgHa mice, each purchased from The Jackson Laboratory (stock no. 002014 and 008341, respectively). CD45.2+ IgHb B cells (10 × 106) purified from Rosa26-CreERT2 ± Prkaa1f/f mice, polyclonal wild-type CD4+ T cells (4 × 106), and CD4+ T cells from OT-II transgenic mice (1 × 106) were transferred into allotype-disparate CD45.1+ IgHa mice preconditioned with sublethal irradiation (split dose of 3.75 Gy × 2) 2 d prior to transfer. These chimeric mice were immunized with NP16-OVA (Biosearch Technologies) emulsified in Imject Alum (100 μg in 100 μl).

Genomic DNA was extracted from sorted cells using cell lysis buffer (1% SDS, 50 mM Tris base, and 10 mM EDTA [pH 8]), and 100 ng genomic DNA was used as a template for Prkaa1 and CreERT2 amplifications using primers previously described (26).

Unless otherwise specified, all mAbs were from BD Pharmingen (San Jose, CA), Life Technologies (Carlsbad, CA), eBioscience (San Diego, CA), or Tonbo Biosciences (San Diego, CA). For detection of GC-phenotype B cells and MphenBC, splenocytes (3 × 106) were stained with anti-B220, -GL7, -Fas, -IgD, and -CD38 and nitrophenol (NP)-allophycocyanin and a dump channel containing anti-CD11b, -CD11c, -F4/80, and -Gr-1 and 7-aminoactinomycin D in 1% BSA and 0.05% sodium azide in PBS. To phenotype different memory B cell subsets, a second panel consisted of anti-B220, -CD38, -CD80, -PD-L2, and -IgG1 and NP-allophycocyanin and the aforementioned dump channel with the addition of anti-IgD, -CD4, -CD8, and -GL7. In analyses of transfer experiments, allotype-specific Abs were used to distinguish donor (CD45.2) and recipient (CD45.1) B cells as described (27). For flow analyses of mitochondria, 1 × 106 to 3 × 106 cells were washed in PBS and stained with 200 nM MitoTracker Green (Invitrogen, Carlsbad, CA), 50 nM MitoTracker Deep Red (Invitrogen), and Ghost-780 in PBS for 20 min at 37°C, then washed again (1% BSA in PBS) and further stained with anti-B220, -CD138, or -CD38. MitoSOX (5 μM) and BODIPY C-11 581/591 (1.25 μM) staining were performed similarly. For intracellular phospho-flow analysis, cells were fixed with 4% PFA, followed by methanol permeabilization. Peptide-specific, anti–phospho-S6 (Cell Signaling Technologies) was as described (16). Samples were analyzed using an FACSCanto flow cytometer driven by BD FACSDiva software and were processed using FlowJo software (FlowJo, Ashland, OR).

Starting from single-cell suspensions of splenocytes, MphenBC were enriched by depleting IgD+ and Thy1.2+ cells using biotinylated Abs, followed by BD IMag streptavidin particles on an IMag Cell Separation Magnet (BD Biosciences, San Jose, CA). Cells were stained as described for flow cytometry and dump-negative, IgD CD38+, B220+ cells were sorted into 10% FBS in PBS for downstream applications.

For detection of circulating NP-binding Ab of all- and high-affinity after immunization with NP-KLH in alum adjuvant, serial dilutions of sera were added to NP24-BSA (Biosearch Technologies)–coated (0.1 μg/well), 96-well plates (Costar, Washington, DC) and incubated overnight at 4°C, followed by incubation with either HRP-conjugated anti-IgM or -IgG1 (SouthernBiotech) or biotinylated anti-IgG1a or anti-IgG1b (BD Biosciences) for transfer experiments, followed by streptavidin–HRP (R&D Systems, Minneapolis, MN). The plates were developed using Ultra TMB Substrate (Thermo Fisher Scientific), and optical densities at 450 nm were measured. To compile results across biologically independent experiments, optical densities within the linear range of serially diluted sera were combined. Using OD values generated from a dilution in the linear range of the curve, “recall effects” were calculated as described (28) using ELISA data from each mouse and subtracting the OD value from its serum collected the day before rechallenge from the OD value of the serum 1 wk post-rechallenge. For detection of secreted Ab after in vitro studies, supernatants from cultured cells were added to anti-Ig(H+L) (SouthernBiotech, Birmingham, AL)–coated, 96-well plates before detection with HRP-conjugated Abs. Normal mouse IgG (Thermo Fisher Scientific) was used as a standard to interpolate concentrations of IgG for tissue culture supernatants.

To detect Ag-specific ASCs after immunization, 96-well, high protein-binding membrane plates (MilliporeSigma, Burlington, MA) were coated with 1 μg/well NP24-BSA. Splenocytes or bone marrow cells (5–20 × 105) were added, and plates were incubated at 37° overnight, followed by incubation with biotinylated anti-IgM, -IgG1, -IgG1a, or -IgG1b Abs prior to incubation with VECTASTAIN ABC Kit (Vector Laboratories, Burlingame, CA) and development using 3-amino-9-ethylcarbzole (Sigma-Aldrich, St. Louis, MO). ASCs were quantified using an ImmunoSpot Analyzer (Cellular Technology, Shaker Heights, OH).

Splenic B cells were purified from Rosa26-CreERT2 mice (Prkaa1+/+ and Prkaa1f/f) by negative selection using biotinylated anti-CD43, -Thy1.2, and -F4/80 (>85% CD19+), followed by streptavidin particles and IMag Cell Separation Magnet (BD Biosciences). To induce plasma cell differentiation, B cells were seeded at 5 × 105 per ml and treated with 5 μg/ml LPS (Sigma-Aldrich), 10 ng/ml BAFF (AdipoGen, San Diego, CA), 10 ng/ml IL-4 (PeproTech, Rocky Hill, NJ), 5 ng/ml IL-5 (PeproTech), and 50 nM 4-hydroxytamoxifen (4-OHT) (Sigma-Aldrich). Cells were cultured for 2–8 d in RPMI 1640 supplemented with 10% FBS (Peak, Denver, CO), 100 U/ml penicillin (Invitrogen), 100 μg/ml streptomycin (Invitrogen), 3 mM l-glutamine (Invitrogen), and 0.1 mM 2-ME (Sigma-Aldrich). B cells were also expanded on the NB21 feeder line as previously described (29, 30). Every 3 d, supernatants were frozen for further analysis, and the expanded B cells were reseeded on fresh NB21 feeder cells in new media and 4-OHT. For spontaneous Ab secretion, day 8 LPS cultures or day 9 NB21.2D9 cultures were subjected to Ficoll spin (Invitrogen) to eliminate dead cells and debris. Cells were then washed, and 5 × 104 cells were seeded in 100 μl of fresh media in a 96-well plate for 8 h. Supernatants were frozen, and levels of secreted IgG1 were determined by ELISA.

Unless otherwise indicated, all immunoblots depict relative protein from whole B cell extracts derived from splenocytes of Rosa26-CreERT2 mice (Prkaa1+/+ or Prkaa1f/f), followed by a 2-d culture with LPS, BAFF, and 4-OHT at 5 × 106/ml. For immunoblotting for AMPK targets after glucose starvation, 2-d LPS blasts were washed and reseeded in glucose-free RPMI 1640 (Invitrogen) supplemented with 10% dialyzed HyClone FBS (Thermo Fisher Scientific) for specified amount of time. Cells were washed twice in cold PBS and lysed in radioimmunoprecipitation assay buffer (catalog no. 0278; Sigma-Aldrich) in the presence of phosphatase inhibitor (Thermo Fisher Scientific) and a protease inhibitor mixture (Sigma-Aldrich). Twenty to one hundred micrograms of cell lysates were resolved by SDS-PAGE, transferred to polyvinylidene difluoride membranes, and immunoblotted for phospho-AMPKα1 (T172), AMPKα, phospho-S6 (S235/236), total S6, phospho-ACC (S79), total ACC, phospho–Unc-51–like kinase (ULK1) (S317), total ULK1, phospho-4E-BP1, total 4E-BP1, and/or LC3 using monoclonal rabbit Abs purchased from Cell Signaling Technology (Danvers, MA). Actin (Santa Cruz Biotechnology, Dallas, TX) was detected on all blots as a loading control. Immunoblots were visualized using Odyssey Imaging System (LI-COR Biosciences) after incubation with secondary reagents, anti-rabbit IgG-680, or anti-mouse IgG-800 (Invitrogen).

Oxygen consumption rate (OCR) and extracellular acidification rate were measured using an XFe96 extracellular flux analyzer (Seahorse Bioscience). Briefly, 5 × 105 2-d LPS- and BAFF-activated B cells were seeded per well of a Cell-Tak (5 μg/ml; Corning) coated plate. Glycolytic and mitochondrial stress tests were performed as previously described (16, 31). Maximum respiration and spare respiratory capacity were calculated using formulas derived from the Seahorse platform.

LPS blasts (5 × 105 cells from 48-h cultures) in 0.5 ml were seeded on poly-d-lysine–coated coverslips in a 24-well plate and stained with 100 nM MitoTracker Deep Red for 20 min at 37° before centrifugation to ensure cellular adherence to coverslips. Coverslips were incubated overnight with anti-B220 or CD138-PE and either anti-LC3, anti-ULK1, or anti-Lamp1 using rabbit mAbs (Cell Signaling Technology) after methanol fixation and blocking in 1% BSA in PBS with Tween 20. Coverslips then were mounted onto slides using ProLong Gold Antifade Reagent (Invitrogen) after incubation with secondary Ab anti-rabbit IgG 488 (Invitrogen) to visualize rabbit Abs using an Olympus FV-1000 fluorescent confocal microscope. LC3-puncta were assessed using ImageJ. Colocalization of lysosomes and mitochondria as an indicator of mitophagy was determined using Just Another Colocalization Plugin in ImageJ. Manders coefficient represents the percentage of mitochondrial pixels (blue channel) that overlay Lamp1 or ULK1 pixels (green channel), in which 0 = no colocalization, and 1 = 100% colocalization.

After 7 d of culture with LPS, BAFF, IL-5, IL-4, and 4-OHT as above, viable cells were recovered after Ficoll step gradient centrifugation, rinsed, and recounted. Equal numbers of cells were then pulsed for 1 h with 2 μCi [3H]-leucine (60 Ci/mmol; Moravek, Brea, CA) per 2 × 106 cells/ml arginine-, lysine-, and leucine-deficient RPMI 1640 (Sigma-Aldrich) supplemented back with complete RPMI 1640 levels of l-arginine (1.149 mM; Sigma-Aldrich), l-lysine (0.219 mM; Sigma-Aldrich), and 10% of complete RPMI 1640 levels of l-leucine (0.038 mM; Sigma-Aldrich). Media also contained 10% dialyzed HyClone FBS, 25 mM HEPES (Invitrogen), and phenol red (Sigma-Aldrich). After pulsing cells, 2 × 106 cells were lysed in radioimmunoprecipitation assay buffer in the presence of protease and phosphatase inhibitors, and supernatants were frozen for the zero-chase time point. Remaining cells were washed in PBS and resuspended in the aforementioned media but with 100% complete RPMI 1640 levels of l-leucine (0.382 mM) at 1 × 106 cells per ml. At each specified chase time point, 2 × 106 cells were lysed, and the supernatant collected. Samples were stored at −20°. Secreted and intracellular IgG Abs were purified from supernatant and lysate samples, respectively, using protein G agarose beads (Santa Cruz Biotechnology). Precipitates were subjected to SDS-PAGE in reducing conditions, transferred to polyvinylidene difluoride membranes, and rocked for 30 min in 2 M sodium salicylate (Sigma-Aldrich) prior to fluorography at −80° for 1–14 d as described previously (32). Molar amounts of [3H]-leucine incorporated into Ab were calculated after membranes were subjected to liquid scintillation counting using a Beckman Coulter LS 6500.

We set out to test the role of AMPK on B cell function over the course of an immune response. Previous studies have provided evidence that AMPKα1, encoded by Prkaa1, is the only isoform of the essential catalytic subunit of AMPK expressed in B cells (21, 22, 33). Thus, to generate mice with a conditional B cell–specific deletion of AMPK, we crossed Prkaa1 floxed mice to transgenic animals expressing a hydroxytamoxifen-inducible Cre recombinase under the control of the huCD20 promoter (huCD20-CreERT2). This B lineage–restricted promoter is active from the pre-B to mature B cell stage (34). AMPKα expression was undetectable in LPS-activated B lymphoblasts from tamoxifen-injected Prkaa1f/f huCD20-CreERT2 mice when compared with Prkaa1+/+ huCD20-CreERT2 controls (Fig. 1A). LPS-activated Prkaa1Δ/Δ B cells failed to phosphorylate AMPKα target ACCS79 even in the absence of glucose indicating a loss of AMPK function in Prkaa1-deleted cells (Supplemental Fig. 1). Because there was no detectable band in Prkaa1-deleted cells with the AMPKα Ab, which detects both AMPKα1 and -α2 catalytic isoforms, we conclude that only AMPKα1 is substantially expressed in B cells and that loss AMPKα1 did not lead to compensatory induction of AMPKα2.

FIGURE 1.

Inactivation of Prkaa1 in the B lineage leads to elevated primary Ab responses and initial MphenBC population. (A) Immunoblot for AMPKα (1, 2) and actin expressed in LPS-activated B cells purified after in vivo tamoxifen regimen. Data representative of immunoblots from n = 3 versus three mice. (B) Schematic of immunization strategy using mice harboring a tamoxifen-inducible Cre expressed specifically in the B lineage. (C) Total number of splenic NP+ GC B cells (B220+ IgD Fas+ GL7+ NP+). (D and E) Total and CD80neg PD-L2neg splenic NP+ MphenBC (B220+ IgD GL7- CD38+ NP+). (F) Total splenic NP-specific IgM- and IgG1-secreting cells detected per 1 × 106 plated by ELISpot. (C–F) Each circle (controls, filled circle [•]; Prkaa1Δ/Δ, open circle [○]) represents sample from one mouse with mean ± SEM also displayed. (G and H) Circulating NP-specific IgM and IgG1 after immunization strategy as described in (A). Data represent mean ± SE from four independent experiments with at least n = 9 Prkaa1+/+ versus n = 9 Prkaa1Δ/Δ. *p < 0.05, **p < 0.01, determined by Mann–Whitney U nonparametric t test or ANOVA when appropriate.

FIGURE 1.

Inactivation of Prkaa1 in the B lineage leads to elevated primary Ab responses and initial MphenBC population. (A) Immunoblot for AMPKα (1, 2) and actin expressed in LPS-activated B cells purified after in vivo tamoxifen regimen. Data representative of immunoblots from n = 3 versus three mice. (B) Schematic of immunization strategy using mice harboring a tamoxifen-inducible Cre expressed specifically in the B lineage. (C) Total number of splenic NP+ GC B cells (B220+ IgD Fas+ GL7+ NP+). (D and E) Total and CD80neg PD-L2neg splenic NP+ MphenBC (B220+ IgD GL7- CD38+ NP+). (F) Total splenic NP-specific IgM- and IgG1-secreting cells detected per 1 × 106 plated by ELISpot. (C–F) Each circle (controls, filled circle [•]; Prkaa1Δ/Δ, open circle [○]) represents sample from one mouse with mean ± SEM also displayed. (G and H) Circulating NP-specific IgM and IgG1 after immunization strategy as described in (A). Data represent mean ± SE from four independent experiments with at least n = 9 Prkaa1+/+ versus n = 9 Prkaa1Δ/Δ. *p < 0.05, **p < 0.01, determined by Mann–Whitney U nonparametric t test or ANOVA when appropriate.

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A prior study provided evidence that a B cell–specific deletion of AMPKα1 driven by CD19-Cre had no effect on GC and no substantial change in early plasma cell generation or initial levels of circulating Ab 2 wk after NP-CGG immunization (22). However, the impact of AMPKα1 on other outcomes of GC, including the generation of memory and long-lived plasma cells remained unexamined. We assessed the effect of B cell–intrinsic AMPKα1 on the generation of MphenBC and ASCs, the latter of which peak in formation during late GC at least 3 wk after exposure to Ag (35). To do so, we immunized tamoxifen-treated huCD20-CreERT2 mice (Prkaa1f/f or Prkaa1+/+) with NP-KLH, followed by boosting after 3 wk and analyses of NP-specific humoral responses 1 wk after the booster immunization (Fig. 1B). Consistent with a prior study (22), we observed no difference in NP-specific GC B cells (B220+ NP+ IgDneg GL7+ Fas+) for AMPKα1-deficient B cells (Fig. 1C). However, induced loss of AMPKα1 from mature B cells led to almost a 2-fold increase in NP-specific MphenBC (B220hi NP+ IgDneg GL7neg CD38+) (Fig. 1D). We further examined whether AMPKα1 differentially supported the generation of different MphenBC subsets distinguished by PD-L2 and CD80 expression (4, 36, 37). The double-negative (CD80neg PD-L2neg) MphenBC population, which makes up >85% of total MphenBC are >90% are IgM+ (37). Hapten-binding CD80neg PD-L2neg MphenBC were increased in the absence of AMPKα1 to a similar extent as NP-specific MphenBC (Fig. 1E, Supplemental Fig. 1B). Double-positive Ag-specific MphenBC, which make up ∼5% of total MphenBC, were unaffected in AMPKα1-null mice (Supplemental Fig. 1C). Although we cannot exclude a contribution of renewed proliferation of MphenBC after the boost, we infer that AMPK dampened the initial MphenBC population size, particularly CD80neg PD-L2neg MphenBC, after a short-term prime/boost immunization strategy. Additionally, the frequencies of splenic (Fig. 1F) and bone marrow (Supplemental Fig. 2A) anti-NP IgM- and IgG1-secreting ASCs were unaffected by the loss of AMPKα1 from the B lineage. Despite similar numbers Ag-specific IgM and IgG1 ASCs, there was a substantial increase in circulating anti-NP IgM and IgG1 as early as 2 wk after the initial immunization in the absence of AMPK (Fig. 1G, 1H). The elevated Ag-specific IgG1 concentrations in the sera were maintained several weeks after immunization despite the apparently normal numbers of anti-NP IgG1 ASCs in the marrow (Supplemental Fig. 2B). Collectively, these data indicate that AMPK restrained both the initial CD80neg PD-L2neg MphenBC population and primary Ab production in vivo without an observable increase in the Ag-specific ASCs or GC B cells derived from AMPKα1-deficient B cells.

Several models, none mutually exclusive of another, could account for the apparent paradox of persistently higher Ab concentrations in the absence of any increase in ASCs derived from AMPKα1-deficient B cells. First, loss of AMPKα1 in B cells may lead to increased rates of Ab production per ASC. Second, loss of AMPKα1 from B cells might lead to more formation of plasma cells that generate higher circulating Ab but die before time of analysis. To explore these possibilities, we used in vitro cultures to test the effect of AMPKα1 on plasma cell differentiation and Ab production. For these in vitro studies, B cells purified from tamoxifen-injected Rosa26-CreERT2 mice (Prkaa1f/f or Prkaa1+/+) were cultured with IL-4 on NB-21.2D9 feeder cells described (29, 30). When CD138 and B220 expression were analyzed every 3 d for 9 d of coculture with the BAFF, CD40L, and IL-21–expressing feeder cells, no differences were observed in the number of plasma cells (B220lo CD138+) (Fig. 2A), suggesting that AMPKα1 is dispensable for plasma cell differentiation. To test the effect of B cell AMPKα1 on Ab production throughout the coculture, IgG1 was measured in supernatants. Despite similar numbers of plasma cells, cultures with AMPKα1-deficient B cells accumulated 3-fold higher concentrations of Ab in their supernatants from day 6 to 9 (Fig. 2B). Thus, consistent with the in vivo serologies, loss of AMPKα1 from B cells led to an increase in Ab in the supernatant despite comparable plasma cell numbers. These in vitro results indicated that the increase in Ab in the supernatant observed from AMPKα1-deficient B cells was due to an increase in Ab production on a per cell basis.

FIGURE 2.

Loss of AMPKα1 has no defect on plasma cell differentiation but leads to increased Ab synthesis per cell. (A) Representative flow plots of CD138 versus B220 expression during plasma cell differentiation after B cells from tamoxifen-treated Rosa26-ERT2Cre mice (Prkaa1+/+ or Prkaa1f/f) were cocultured on NB-21.2D9 feeder cells in the presence of 4-OHT and IL-4 (left panel). Total number (mean ± SEM) of plasma cells (CD138+B220lo) throughout coculture (right). (B) Mean (± SEM) concentrations of IgG1 detected in supernatants collected from days 3, 6, and 9 of coculture. (C) Representative wells and quantification of ELISpot analysis depicting numbers of IgG1-secreting cells per 500 plated cells. (D) Mean spot size of IgG1 ASCs after 9 d of coculture. (E) Relative levels of IgG1 detected in the supernatant 8 h after plating 5 × 104/100 μl day 9 cocultured cells. Data normalized to wild-type controls. Data represent mean ± SE from at least two independent experiments with n = 6 Prkaa1+/+ versus n = 4 or 5 Prkaa1Δ/Δ mice. (F) Protein G precipitation of intracellular and secreted IgG1 from 7-d LPS cultures after 1 h of labeling with [3H]-leucine and the indicated chase times. (G) [3H]-leucine incorporation into IgG1 collected from lysates and supernatants at indicated chase times. (H) Ratio of [3H]-leucine incorporation in the supernatant to incorporation in the lysate, derived from (G), as an indicator of secretion efficiency. Data are representative of three independent experiments using n = 3 Prkaa1+/+ versus n = 3 Prkaa1Δ/Δ mice. (C–H) Each circle (controls, filled circle [•]; Prkaa1Δ/Δ, open circle [○]) represents sample from one mouse, with mean ± SEM also displayed. **p < 0.01, ***p < 0.001, ****p < 0.0001, determined by Mann–Whitney U nonparametric t test or ANOVA when appropriate.

FIGURE 2.

Loss of AMPKα1 has no defect on plasma cell differentiation but leads to increased Ab synthesis per cell. (A) Representative flow plots of CD138 versus B220 expression during plasma cell differentiation after B cells from tamoxifen-treated Rosa26-ERT2Cre mice (Prkaa1+/+ or Prkaa1f/f) were cocultured on NB-21.2D9 feeder cells in the presence of 4-OHT and IL-4 (left panel). Total number (mean ± SEM) of plasma cells (CD138+B220lo) throughout coculture (right). (B) Mean (± SEM) concentrations of IgG1 detected in supernatants collected from days 3, 6, and 9 of coculture. (C) Representative wells and quantification of ELISpot analysis depicting numbers of IgG1-secreting cells per 500 plated cells. (D) Mean spot size of IgG1 ASCs after 9 d of coculture. (E) Relative levels of IgG1 detected in the supernatant 8 h after plating 5 × 104/100 μl day 9 cocultured cells. Data normalized to wild-type controls. Data represent mean ± SE from at least two independent experiments with n = 6 Prkaa1+/+ versus n = 4 or 5 Prkaa1Δ/Δ mice. (F) Protein G precipitation of intracellular and secreted IgG1 from 7-d LPS cultures after 1 h of labeling with [3H]-leucine and the indicated chase times. (G) [3H]-leucine incorporation into IgG1 collected from lysates and supernatants at indicated chase times. (H) Ratio of [3H]-leucine incorporation in the supernatant to incorporation in the lysate, derived from (G), as an indicator of secretion efficiency. Data are representative of three independent experiments using n = 3 Prkaa1+/+ versus n = 3 Prkaa1Δ/Δ mice. (C–H) Each circle (controls, filled circle [•]; Prkaa1Δ/Δ, open circle [○]) represents sample from one mouse, with mean ± SEM also displayed. **p < 0.01, ***p < 0.001, ****p < 0.0001, determined by Mann–Whitney U nonparametric t test or ANOVA when appropriate.

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We next characterized differentiated cells at day 9 of coculture, when the greatest difference in Ab production with the loss of AMPKα1 was observed. To test if similar frequencies of B220lo CD138+ cells translated to similar numbers of functional ASCs, we performed ELISpot analyses. These assays detected ∼40 IgG1-secreting cells per 500 plated differentiated cells for both AMPKα1-deficient and -sufficient B cells (Fig. 2C). Despite similar numbers of IgG1-secreting cells after 9 d of coculture with the loss of AMPKα1, the mean spot size for AMPKα1-deficient plasma cells was increased (Fig. 2D). This finding suggested that the amount of Ab secreted per cell was greater with the loss of AMPKα1. To test this rigorously, we measured the effect of AMPKα1 on short-term (8 h) Ab secretion after replating equal numbers of in vitro–differentiated cells in fresh media. IgG1 detected in the supernatant from AMPKα1-deficient cells was ∼1.5-fold greater than from controls (Fig. 2E). Collectively, data indicate that AMPKα1 in B cells neither hinders nor promotes plasma cell generation but attenuates the amount of Ab produced per plasma cell.

We next examined whether the increase in Ab production per cell from AMPKα1-deficient B cells was due to an increase in Ig synthesis and/or enhanced activity of the secretory pathway for Ig. To obtain enough plasma cells for downstream biochemical assays, we activated and cultured wild-type and AMPKα1-deficient B cells using LPS. Similar to the coculture system, the frequency of plasma cells generated by LPS, BAFF, IL-4, and IL-5 were unaffected with the loss of AMPKα1 (Supplemental Fig. 2C), but IgG1 detected in the supernatant from AMPKα1-deficient cells were enhanced 2-fold (Supplemental Fig. 2D).

To distinguish the role of AMPKα1 on rates of Ab synthesis and/or Ig secretion, we performed [3H]-leucine pulse-chase analyses after replating equal numbers of LPS-differentiated cells. AMPKα1-deficient cells exhibited increased intracellular and secreted [3H]-leucine–labeled Ig (Fig. 2F, 2G). Specifically, AMPKα1-deficient cultures had ∼6.5 fmol [3H]-leucine incorporated in Ab after 1h of labeling compared with ∼3.5 fmol of wild-type controls (Fig. 2G). The similar ratio of secreted to intracellular [3H]-Ab after labeling was most consistent with increased synthesis as the basis for higher IgG1 production (Fig. 2H). Together, these data demonstrate that AMPKα1 in B cells restrains rates of Ab synthesis in plasma cells.

AMPKα1 inhibits mTORC1 activity both indirectly through the activation of TSC2 and directly by an inhibitory phosphorylation of Raptor, an essential component of mTORC1 (38). Loss of Raptor in B cells led to poor Ab responses and inefficient generation of memory B cells in vivo (16). To evaluate mTORC1 activity in AMPKα1-deficient B cells, we performed immunoblots for target phospho-proteins of mTORC1 using extracts of B lymphoblasts. Consistent with the canonical model, loss of AMPKα1 led to elevated expression of downstream mTORC1 targets, phospho-S6S235/236, and phospho-4E-BP1T37/46 (Fig. 3A, 3B). Similarly, splenic MphenBC from Prkaa1f/f huCD20-CreERT2 mice had elevated levels of phospho-S6 compared with wild-type controls after undergoing the immunization strategy illustrated in Fig. 1B (Fig. 3C, top panel). Enhanced phospho-S6 in AMPKα1-null plasma cells was also observed in the bone marrow (Fig. 3C, bottom panel). Because mTORC1 promotes protein synthesis, the elevated levels of Ab production observed with AMPKα1-deficient plasma cells are consistent with the increase in mTORC1 activity.

FIGURE 3.

Increased mTORC1 activity in the absence of AMPKα1 in the B lineage. (A and B) Immunoblot of mTORC1 targets pS6S235/236 (A) and p4E-BP1T37/46 (B) after 2 d of LPS and BAFF activation. Data represent three independent experiments or quantified in the right panels as the ratio of the indicated phospho-protein to signal for the holoprotein. The p values were determined by paired t test. (C) Representative plot and quantification of the mean fluorescence intensity of phospho-S6 in splenic MphenBC (B220+IgDGL7CD38+) and bone marrow plasma cells (CD138+TACI+). Data represent two independent experiments with n = 6 versus six mice. The p values were determined by Mann–Whitney U nonparametric t test.

FIGURE 3.

Increased mTORC1 activity in the absence of AMPKα1 in the B lineage. (A and B) Immunoblot of mTORC1 targets pS6S235/236 (A) and p4E-BP1T37/46 (B) after 2 d of LPS and BAFF activation. Data represent three independent experiments or quantified in the right panels as the ratio of the indicated phospho-protein to signal for the holoprotein. The p values were determined by paired t test. (C) Representative plot and quantification of the mean fluorescence intensity of phospho-S6 in splenic MphenBC (B220+IgDGL7CD38+) and bone marrow plasma cells (CD138+TACI+). Data represent two independent experiments with n = 6 versus six mice. The p values were determined by Mann–Whitney U nonparametric t test.

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Early after a prime/boost primary Ab response, B lineage–intrinsic AMPKα1 limited the MphenBC population (Fig. 1D, 1E) and Ab synthesis (Fig. 1G, 1H). Because mTORC1 hyperactivity in TSC1-deficient T lineage cells undermined their longevity (39), we assessed if AMPKα1 affects the persistence of MphenBC in vivo, bearing in mind that mTORC1 activity in MphenBC is decreased compared with the overall pool of GC B cells as indicated by flow cytometric measurements of phospho-S6 in these two populations (Supplemental Fig. 2E). Because of technical issues relating to new B cell production after tamoxifen treatments in the huCD20-CreERT2 mice, we used the potent B lineage–specific mb1-Cre transgene to drive excision of Prkaa1 conditional alleles (25). This mode of excision, which starts as early as the pro–B cell stage, is constitutive but tamoxifen independent. Lack of AMPKα1 throughout B lymphoid ontogeny had no discernable effect on the preimmune B cell populations of mb1-Cre Prkaa1f/f mice (Supplemental Fig. 3A, 3B). Similar to primary response data observed in huCD20-CreERT2 mice, Prkaa1f/f mb1-Cre mice exhibited normal frequencies of NP-specific GC B cells 1 wk after immunization compared with wild-type controls (Supplemental Fig. 3C). To test the effect of AMPK on the longevity of the MphenBC population, we immunized mb1-Cre mice (Prkaa1f/f and Prkaa1+/+) with NP-KLH, analogous to Fig. 1B–F, except mice were harvested 8 wk after the boost (Fig. 4A) instead of 1 wk (Fig. 1). In sharp contrast to the increase in total and CD80neg PD-L2neg Ag-specific MphenBC populations 1 wk postboost (Fig. 1D), the frequencies of these MphenBC populations were reduced 8 wk postboost when B lineage were AMPKα1 deficient (Fig. 4B, 4C). Together with our data at the early time point after immunization (Fig. 1D), we infer that although AMPKα1 limits MphenBC generation/expansion during an early phase of a primary response, while GC are active, it enhances the long-term persistence of memory B cells defined by standard phenotypic criteria (i.e., MphenBC).

FIGURE 4.

AMPKα1 supports the long-term persistence of Ag-specific MphenBC and recall humoral response. (A) Schematic of immunization strategy with a hapten carrier using mice harboring mb1-Cre, a tamoxifen-independent Cre recombinase expressed specifically in the B lineage. (B) Frequency of MphenBC that are NP specific. (C) Frequency of NP-specific PD-L2negCD80neg within the MphenBC gate. Data represent three independent experiments with 10 versus 10 mice. The p values were determined by Mann–Whitney U nonparametric t test. (D) Schematic of immunization strategy to assess humoral recall. (E) Quantitation of the magnitudes of increase in anti-NP IgM and (F) anti-NP IgG1 in sera 1 wk after recall immunization relative to prerecall concentrations. Shown are the calculated recall effects as defined in (28) and the 2Materials and Methods. (G) Total numbers of splenic CD138+TACI+ cells and (H) NP-specific GC B cells 1 wk after recall challenge.

FIGURE 4.

AMPKα1 supports the long-term persistence of Ag-specific MphenBC and recall humoral response. (A) Schematic of immunization strategy with a hapten carrier using mice harboring mb1-Cre, a tamoxifen-independent Cre recombinase expressed specifically in the B lineage. (B) Frequency of MphenBC that are NP specific. (C) Frequency of NP-specific PD-L2negCD80neg within the MphenBC gate. Data represent three independent experiments with 10 versus 10 mice. The p values were determined by Mann–Whitney U nonparametric t test. (D) Schematic of immunization strategy to assess humoral recall. (E) Quantitation of the magnitudes of increase in anti-NP IgM and (F) anti-NP IgG1 in sera 1 wk after recall immunization relative to prerecall concentrations. Shown are the calculated recall effects as defined in (28) and the 2Materials and Methods. (G) Total numbers of splenic CD138+TACI+ cells and (H) NP-specific GC B cells 1 wk after recall challenge.

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We next sought to test whether the decline in MphenBC in the absence of AMPKα1 had consequences on humoral recall (i.e., memory function). Upon subsequent encounter with Ag, memory B cells rapidly differentiate into ASCs or reenter GC to generate faster and more robust humoral responses than at the initial encounter. Accordingly, we rechallenged mb1-Cre mice (Prkaa1f/f versus Prkaa1+/+) 14 wk after a primary immunization (Fig. 4D). To assess the strength of recall responses Prkaa1-null MphenBC, we measured circulating anti-NP IgM and anti-NP IgG1 in sera collected both immediately prior to and 1 wk after rechallenge, thereby allowing for the possible persistence of higher Ab levels in the absence of AMPKα1 (Fig. 1H). For each mouse, then, a recall effect (mnestic induction of increased Ab concentration) was calculated (28). Anti-NP IgM and IgG1 increased substantially less in response to the recall immunization of Prkaa1f/f mb1-Cre mice than mb1-Cre controls (Fig. 4E, 4F). These data indicate that in contrast to the primary response (Fig. 1), recall Ab responses were weaker when B cells lacked AMPKα1. Consistent with the observed decrease in recall-induced Ab, the CD138+ TACI+ splenic plasmablast/plasma cell populations (40) also were attenuated in mice harboring AMPKα1-deficient B cells (Fig. 4G). In contrast to the primary response, Ag-specific GC B cells, a fate of reactivated CD80neg PD-L2neg MphenBC (4, 36), were also diminished in rechallenged AMPKα1-deficient mice. We infer from these data that AMPKα1 not only supports the longevity of MphenBC but also maintains the functional capacity of recall humoral responses.

Mice in the previous model (Prkaa1f/f mb1-Cre) harbor B cells that are deficient in AMPKα1 from very early in B lineage development so that a defect of programming during the primary response might contribute to the finding of weaker recall. To test the role of AMPKα1 after primary immunity, we used adoptive transfers with a tamoxifen-inducible system in which Prkaa1 can be deleted after a normal primary response (Fig. 5A). Naive B cells (CD45.2+ IgHb allotype) from Rosa26-CreERT2 mice (Prkaa1f/f or Prkaa1+/+) were transferred into sublethally irradiated CD45.1+ IgHa allotype–disparate recipients along with Th cells (a mix of CD4+ T cells from wild-type and OT-II transgenic mice). Recipients were then immunized with NP-OVA and boosted (as in Fig. 1A) to generate normal primary responses. Tamoxifen injections to induce deletion of Prkaa1 in the donor B cell population, including MphenBC, were deferred to week 9. To assess recall responses, mice were rechallenged with NP-OVA several months later and harvested 1 wk after the recall immunization (Fig. 5A). PCR products from flow-sorted donor CD45.2+ MphenBC at the time of harvest documented effective deletion of Prkaa1 in the cells from mice that received Prkaa1f/f Rosa26-CreERT2 donor B cells (Fig. 5B). Using allotype-specific detection reagents, donor and recipient anti-NP IgG1 were measured in the sera before and after recall challenge. Compared with mice receiving wild-type CreERT2 B cells, mice that received CreERT2Prkaa1f/f B cells had weaker anti-NP IgG1b recall responses after rechallenge (Fig. 5C). In contrast, an internal control of recipient-derived a-allotype Ab found that levels of anti-NP IgG1a were similar regardless of the donor cell source. Consistent with these serologies after rechallenge, anti-NP IgG1b–secreting cells and donor-derived CD45.2+ CD138+ TACI+ plasmablasts and plasma cells in the spleen were diminished in mice that received B cells from which AMPKα1 was depleted (Fig. 5D, 5E). AMPKα1-deficient, donor-derived GC B cells were also decreased compared with wild-type donor GC B cells (Fig. 5F). These data support the conclusion that after a normal primary humoral response is established, the recall capacity of B cells is enhanced by AMPKα1.

FIGURE 5.

Loss of AMPKα1 after primary immunity impairs subsequent recall function. (A) Experimental design to evaluate the effect of AMPKα1 in MphenBC after a normal primary response. CD4+ T cells and CD45.2+ IgHb B cells from Rosa26- CreERT2 (Prkaa1f/f or Prkaa1+/+) mice were transferred into naive, wild-type, allotype-disparate CD45.1+ IgHa mice prior to immunization. Deletion of Prkaa1 in MphenBC was induced by tamoxifen injections 9 wk after the initial immunization. Mice were rechallenged 18 wk thereafter and harvested 1 wk after the recall immunization. (B) Prkaa1 and Cre PCR of DNA from purified splenic CD45.1+ or CD45.2+ MphenBC (B220+ IgD GL7 CD38+). (C) Circulating anti-NP IgG1a (recipient derived) and anti-NP IgG1b (donor derived) immediately before and 1 wk post-rechallenge. (D) Representative wells and quantification of ELISpot analyses determining numbers of splenic NP-specific IgG1b ASCs at harvest. (E) Numbers of donor CD45.2+ CD138+ TACI+ plasma cells in the spleen after rechallenge. (F) Left, Representative flow plots depicting frequency of splenic NP-specific GC B cells that are recipient derived (CD45.1+) versus donor derived (CD45.2+) after rechallenge. Right, Total number of donor-derived, NP-specific GC B cells in the spleen after rechallenge.

FIGURE 5.

Loss of AMPKα1 after primary immunity impairs subsequent recall function. (A) Experimental design to evaluate the effect of AMPKα1 in MphenBC after a normal primary response. CD4+ T cells and CD45.2+ IgHb B cells from Rosa26- CreERT2 (Prkaa1f/f or Prkaa1+/+) mice were transferred into naive, wild-type, allotype-disparate CD45.1+ IgHa mice prior to immunization. Deletion of Prkaa1 in MphenBC was induced by tamoxifen injections 9 wk after the initial immunization. Mice were rechallenged 18 wk thereafter and harvested 1 wk after the recall immunization. (B) Prkaa1 and Cre PCR of DNA from purified splenic CD45.1+ or CD45.2+ MphenBC (B220+ IgD GL7 CD38+). (C) Circulating anti-NP IgG1a (recipient derived) and anti-NP IgG1b (donor derived) immediately before and 1 wk post-rechallenge. (D) Representative wells and quantification of ELISpot analyses determining numbers of splenic NP-specific IgG1b ASCs at harvest. (E) Numbers of donor CD45.2+ CD138+ TACI+ plasma cells in the spleen after rechallenge. (F) Left, Representative flow plots depicting frequency of splenic NP-specific GC B cells that are recipient derived (CD45.1+) versus donor derived (CD45.2+) after rechallenge. Right, Total number of donor-derived, NP-specific GC B cells in the spleen after rechallenge.

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AMPK regulates many aspects of intracellular metabolism, including supporting autophagy, the degradation, and recycling of cellular components (41). Autophagy appears to support the survival of memory B cells and limits Ab production in favor of sustaining a long-lived plasma cell population (1113). We hypothesized that the decline in the MphenBC population and/or increased Ab synthesis observed in B cell–specific, AMPKα1-deficient mice was due to a defect in autophagy in the B lineage. Surprisingly, the ability of LPS-activated B cells to form LC3-puncta indicative of autophagosome formation after glucose starvation was no different in AMPKα1-deficient B cells compared with controls (Supplemental Fig. 4A, 4B). Furthermore, LC3-I conversion into the faster migrating LC3-II upon glucose starvation was independent of AMPKα1 expression (Supplemental Fig. 4C). Thus, LPS-activated B cells appear able to undergo glucose starvation-induced autophagy by an AMPKα1-independent mechanism. Finally, normal frequencies and sizes of LC3-puncta were visualized in freshly purified bone marrow plasma cells of tamoxifen-treated Prkaa1f/f, Rosa26-CreERT2 mice (Supplemental Fig. 4D). This result further supports that autophagy occurs in the B lineage even in the absence of AMPKα1. Collectively, these data suggest that B lineage cells can induce AMPKα1-independent autophagy, potentially by noncanonical pathways (42).

In light of the role of AMPK as a regulator of intermediary metabolism and mitochondrial function, we next tested metabolic performance of activated B cells that were AMPKα1-sufficient or -deficient. Extracellular flux analyses revealed no changes in any aspect of the extracellular acidification rate with the loss of AMPKα1 (Fig. 6A). In contrast, analyses of the OCR, before and after treatment with different mitochondrial stressors, determined that mitochondrial oxidative phosphorylation was impaired in AMPKα1-deficient B cells (Fig. 6B). Basal respiration, represented by the OCR values before the addition of ATP synthase V inhibitor oligomycin, was not altered by the loss of AMPKα1 (Fig. 6B, left panel). However, loss of AMPKα1 led to defects in maximal and spare respiratory capacity (Fig. 6B, middle and right panels). The OCR was specific to the electron transport chain, as the OCR levels were dependent on complex I and III inhibitors rotenone and antimycin A. Taken together, these data indicate that AMPKα1 in activated B cells promotes the establishment or maintenance of optimal respiratory function of mitochondria.

FIGURE 6.

Loss of AMPKα1 in B cells leads to decreased mitochondrial function and defects in mitophagy. (A) Glycolytic stress test after B cells from tamoxifen-treated Rosa26-ERT2Cre mice (Prkaa1+/+ or Prkaa1f/f) were activated for 2 d with BAFF and LPS in the presence of 4-OHT. (B) Mitochondrial stress test after B cells from tamoxifen-treated Rosa26-ERT2Cre mice ± Prkaa1f/f were activated for 2 d with BAFF and LPS in the presence of 4-OHT (left). Maximal respiration and spare respiratory capacity calculated by Seahorse report generator (right). Data represent three independent experiments with n = 9 versus eight mice. (C) Expression of pULK1S317, an AMPK target that initiates mitophagy, after glucose deprivation in day 2 LPS-activated cells. Data are representative of three independent experiments. (D) Relative mean fluorescence intensity (MFI) values for MitoTracker Deep Red (left) and MitoTracker Green (right) in the B220+ gate after activation with LPS, BAFF, IL-4, and IL-5 in the presence of 4-OHT. Data represent four independent experiments with 10 versus 11 mice. (E) Quantification of ULK1 and MitoTracker Deep Red colocalization after 2 d activation with LPS and BAFF. (F) Representative immunofluorescence of MitoTracker Deep Red and Lampl colocalization on day 2 LPS-activated cells. (blue, MitoTracker Deep Red; green, Lamp1; red, B220; original magnification ×100). Quantification of Lamp1 and MitoTracker Deep Red colocalization (right). Data are representative of two independent experiments with n = 4 versus three mice and 10 fields per mouse. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, determined by Mann–Whitney U nonparametric t test or ANOVA when appropriate.

FIGURE 6.

Loss of AMPKα1 in B cells leads to decreased mitochondrial function and defects in mitophagy. (A) Glycolytic stress test after B cells from tamoxifen-treated Rosa26-ERT2Cre mice (Prkaa1+/+ or Prkaa1f/f) were activated for 2 d with BAFF and LPS in the presence of 4-OHT. (B) Mitochondrial stress test after B cells from tamoxifen-treated Rosa26-ERT2Cre mice ± Prkaa1f/f were activated for 2 d with BAFF and LPS in the presence of 4-OHT (left). Maximal respiration and spare respiratory capacity calculated by Seahorse report generator (right). Data represent three independent experiments with n = 9 versus eight mice. (C) Expression of pULK1S317, an AMPK target that initiates mitophagy, after glucose deprivation in day 2 LPS-activated cells. Data are representative of three independent experiments. (D) Relative mean fluorescence intensity (MFI) values for MitoTracker Deep Red (left) and MitoTracker Green (right) in the B220+ gate after activation with LPS, BAFF, IL-4, and IL-5 in the presence of 4-OHT. Data represent four independent experiments with 10 versus 11 mice. (E) Quantification of ULK1 and MitoTracker Deep Red colocalization after 2 d activation with LPS and BAFF. (F) Representative immunofluorescence of MitoTracker Deep Red and Lampl colocalization on day 2 LPS-activated cells. (blue, MitoTracker Deep Red; green, Lamp1; red, B220; original magnification ×100). Quantification of Lamp1 and MitoTracker Deep Red colocalization (right). Data are representative of two independent experiments with n = 4 versus three mice and 10 fields per mouse. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, determined by Mann–Whitney U nonparametric t test or ANOVA when appropriate.

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AMPK phosphorylates multiple downstream targets that support different aspects of mitochondrial homeostasis and quality, including mitochondrial biogenesis and mitophagy (i.e., the selective degradation of mitochondria) (43). As we observed no defect in overall autophagy in the mutant B cells, we explored whether AMPK was essential for mitophagy in LPS-activated B cells. ULK1, a substrate of AMPK, is involved in the initiation of canonical autophagy and is essential for mitophagy (4447). LPS-activated wild-type B cells increased phosphorylation of the S317 site of ULK1, a target of AMPK, after 2 h of glucose starvation (Fig. 6C). Induction of ULK1 (S317) phosphorylation due to glucose withdrawal was dependent on AMPKα1 expression. Mitophagy is critical for the removal of damaged or superfluous mitochondria. We reasoned that a defect in mitophagy would lead to the accumulation of dysfunctional mitochondria over time. To assess the effect of AMPKα1 on mitochondria accumulation after in vitro activation, wild-type and mutant B cells were stained with MitoTracker Green and MitoTracker Deep Red, as their fluorescence reports total mitochondrial mass and actively respiring mitochondria, respectively. Mitochondrial mass was comparable regardless of AMPKα1 expression on day 2, but by day 6, AMPKα1-deficient B cells had increased mitochondrial mass in the B220+ gate compared with controls (Fig. 6D, right panel). The increased mitochondrial mass over time was not accompanied with increased mitochondrial activity (Fig. 6D, left panel), suggesting that the relative increased MitoTracker Green staining (day 2 to day 6) represented nonfunctional mitochondria. The evidence is consistent with a model in which AMPKα1 in B cells is important for the clearance of ineffective mitochondria, as characterized for other cell types (48).

Once activated in a manner dependent in part on AMPK-mediated phosphorylation, ULK1 migrates to the mitochondria and triggers a signaling cascade that leads to the recruitment of LC3 machinery and fusion with the lysosome (45, 46). To test if AMPKα1 enhances mitophagy in B cells, we assessed in LPS-activated B cells the association of mitochondria with ULK1 and Lamp1, a lysosomal marker by confocal imaging. Colocalization analyses with wild-type and AMPKα1-deficient B cells revealed that the percentage of ULK1+ mitochondria was decreased in the absence of AMPKα1 (Fig. 5E), suggesting that AMPK plays a role in ULK1 recruitment to mitochondria. As further evidence of a defect in mitophagy, the frequencies of Lamp1+ mitochondria indicative of mitochondrial fusion to the lysosome were diminished by loss of AMPK (Fig. 6F). Collectively, these findings support a model in which AMPKα1 promotes mitochondrial clearance and quality control in B cells, likely due to a nonredundant requirement for ULK1 phosphorylation at S317 and supporting mitochondrial function.

We next tested if the defect in mitochondrial homeostasis applied to the MphenBC population. To obtain sufficient cells, mitochondrial parameters were analyzed by flow cytometry using total rather than NP-specific MphenBC (B220+ GL7neg IgDneg CD38+) 7–10 wk after immunizations of mb1-Cre mice (Prkaa1f/f versus Prkaa1+/+) with NP-KLH (Fig. 7A, left). Actively respiring mitochondria were modestly decreased in the AMPKα1-deficient MphenBC, whereas total mitochondrial mass was unchanged compared with mb1-Cre controls (Fig. 7A, right). The ratio of mitochondrial membrane potential to total mitochondrial mass (Fig. 7A), indicative of mitochondrial quality, was diminished in AMPKα1-deficient MphenBC (Fig. 7B) (49). These data indicate that AMPKα1 supports mitochondrial quality maintenance in MphenBC. We next tested if the decrease in mitophagy observed in LPS blasts (Fig. 6F) was also observed MphenBC immediately ex vivo from mb1-Cre mice (Prkaa1f/f versus Prkaa1+/+). Colocalization analysis revealed that MphenBC from mb1-Cre Prkaa1f/f mice harbored fewer Lamp1+ mitochondria, consistent decreased levels of mitophagy in memory B cells lacking AMPKα1 (Fig. 7C). Levels of mitochondrially derived reactive oxygen species (mtROS) play a key role in many cellular functions, including regulating B cell fate and function (50, 51). Consistent with the defect in quality control, AMPKα1-deficient MphenBC exhibited increased mtROS compared with controls (Fig. 7D). To test if increased mtROS observed in AMPKα1-deficient MphenBC was accompanied with changes in lipid peroxidation, we used BODIPY 581/591 C-11 and found it to be elevated in AMPKα1-deficient MphenBC (Fig. 7E). Collectively, these results suggest that AMPKα1 protects memory B cells against excessive oxidative stress and cell death involving lipid peroxidation by contributing to mitophagic quality control.

FIGURE 7.

AMPKα1 supports mitochondrial quality and limits lipid peroxidation in MphenBC. (A) Left, Representative flow plots of MitoTracker Deep Red versus MitoTracker Green of MphenBC (B220+CD38+ after IgD depletion) from mb1-Cre (Prkaa1f/f versus Prkaa1+/+) mice 11 wk after immunization with a hapten carrier. Right, Quantification of MitoTracker Deep Red and MitoTracker Green mean fluorescence intensity (MFI) values normalized to wild-type. (B) Mitochondrial quality (i.e., the normalized ratio of MitoTracker Deep Red to MitoTracker Green) as an indicator of the proportion of functional mitochondria to the total. (C) Representative images at original magnification ×100 and quantification of the colocalization of MitoTracker Deep Red (blue) and Lamp1 (green) of flow-sorted MphenBC (B220+ GL7 IgD CD38+) from mb1-Cre mice (Prkaa1+/+ or Prkaa1f/f). (D) Representative MitoSOX histogram of MphenBC from mb1-Cre mice 11 wk after immunization with a hapten carrier (left). Quantification of normalized MitoSOX values (right). (E) Representative histogram of BODIPY C-11 (which fluoresces in the FL-1 channel upon lipid oxidation) of MphenBC from mb1-Cre mice 11 wk after immunization (left). Normalized ratio of FITC (FL-1) MFI to PE (FL-2) MFI of MphenBC after staining with BODIPY 581/591 as a measure of lipid peroxidation. Data are representative of three independent experiments with n = 10 versus 10 mice when the average of wild-type values in each experiment was taken and all mice were normalized to that value. The p values were determined by Mann–Whitney U nonparametric t test.

FIGURE 7.

AMPKα1 supports mitochondrial quality and limits lipid peroxidation in MphenBC. (A) Left, Representative flow plots of MitoTracker Deep Red versus MitoTracker Green of MphenBC (B220+CD38+ after IgD depletion) from mb1-Cre (Prkaa1f/f versus Prkaa1+/+) mice 11 wk after immunization with a hapten carrier. Right, Quantification of MitoTracker Deep Red and MitoTracker Green mean fluorescence intensity (MFI) values normalized to wild-type. (B) Mitochondrial quality (i.e., the normalized ratio of MitoTracker Deep Red to MitoTracker Green) as an indicator of the proportion of functional mitochondria to the total. (C) Representative images at original magnification ×100 and quantification of the colocalization of MitoTracker Deep Red (blue) and Lamp1 (green) of flow-sorted MphenBC (B220+ GL7 IgD CD38+) from mb1-Cre mice (Prkaa1+/+ or Prkaa1f/f). (D) Representative MitoSOX histogram of MphenBC from mb1-Cre mice 11 wk after immunization with a hapten carrier (left). Quantification of normalized MitoSOX values (right). (E) Representative histogram of BODIPY C-11 (which fluoresces in the FL-1 channel upon lipid oxidation) of MphenBC from mb1-Cre mice 11 wk after immunization (left). Normalized ratio of FITC (FL-1) MFI to PE (FL-2) MFI of MphenBC after staining with BODIPY 581/591 as a measure of lipid peroxidation. Data are representative of three independent experiments with n = 10 versus 10 mice when the average of wild-type values in each experiment was taken and all mice were normalized to that value. The p values were determined by Mann–Whitney U nonparametric t test.

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The orchestration of intracellular metabolic pathways varies to support developmental and functional needs throughout the lifespan of a B lineage cell (6, 7, 52). Furthermore, as B cells and their progeny migrate through or take residence in distinct tissues, they likely have the capacity to persist in distinct microenvironments with differing nutrient availability (6, 7). Relatively little is known about the metabolic pathways that support the persistence and function of memory B cells and long-lived plasma cells, each of which confers one facet of durable humoral immunity. The energy sensor AMPK regulates multiple aspects of intracellular metabolism. Its overall program favors energy conservation and tilts balance toward catabolism while restraining anabolism in response to declining levels of ATP (14). In this study, we have shown that AMPK supports the longevity of memory B cells essential for robust recall humoral responses and regulates the Ab production of plasma cells both in short- and long-term times after immunization. In particular, the lack of AMPK in B cells led to increased mTORC1 activity as well as initial and long-term Ig synthesis in response to immunization with haptenated carrier. After the initial population increase, AMPK deficiency resulted in declines in the MphenBC population along with the accumulation of dysfunctional mitochondria, decreased mitophagy, increased mtROS, and increased lipid peroxidation. The functional impact of these changes was a reduced strength of recall Ab response after rechallenge. These findings indicate that AMPK protects the maintenance of a memory B cell population and suggest that this function is achieved at least in part through the regulation of mitochondrial turnover. In contrast, AMPK was dispensable for the persistence of bone marrow–resident, long-lived plasma cells but played a critical role in regulating their rate of Ab synthesis.

Immunological memory is a hallmark component of the adaptive immune system whereby long-lived cells of the T and B lineages confer long-lasting protective immunity against reinfection. Although there are some distinctions in the balance of oxidative phosphorylation and glycolysis employed by B and T cells during activation (31) such that B lymphoblasts better couple pyruvate generation to the mitochondria than their T cell counterparts, both subsets undergo clonal expansion and generate long-lived quiescent memory populations (53, 54). Thus, there may be parallels in the metabolic programming between the two lineages. Akin to the role of AMPK in maintaining CD8 memory T cells (18), our data indicate that AMPK promotes the persistence of MphenBC and their function in humoral recall responses. Analogous to reports in the T lineage (18, 19, 55), we found that AMPK antagonizes mTORC1 activity in B cells, a cross-talk that is likely to play a role in regulating rates of protein synthesis. The finding that AMPK promotes longevity and catabolism in both the B and T cell lineages suggests a conserved function of AMPK in these long-lived memory subsets. However, the long-lived plasma cell population provides a second limb of humoral memory (56), and it is not clear if any T cell subset is analogous to this population of terminally differentiated cells that erased most central features of B cell identity (57). Unlike quiescent memory populations, long-lived plasma cells require high rates of protein and glucose metabolism to support the synthesis and glycosylation of Abs (5). Surprisingly, our data indicate that unlike in memory lymphocytes, AMPK is dispensable for maintaining the marrow-resident, long-lived plasma cell population. This finding highlights a stark molecular distinction between memory B cells and bone marrow plasma cells in their metabolic requirements.

In primary humoral responses, mTORC1 is critical for memory B cell generation and Ab production (16, 58). In the current study, we observed enhanced mTORC1 activity in AMPK-deficient ex vivo memory B cells and plasma cells. This measurement corresponded to an increase in the initial number of MphenBC observed 1 wk after a booster immunization as well as enhanced Ab production from plasma cells. Thus, our data suggest that AMPK moderates mTORC1 activity in a manner that maintains a biologically appropriate range of mTORC1 signaling that ultimately leads to appropriate levels of Ab and memory B cell generation. Such “Goldilocks mTOR” may be evolutionally conserved for purposes of maintaining controlled primary humoral responses as it does in T cell differentiation (59). We have uncovered that AMPK plays a part in the fine-tuning of mTORC1 during B cell activation. Elevated mTORC1 signaling and enhanced circulating class-switched Ab were maintained in AMPK-deficient, long-lived bone marrow plasma cells. In light of evidence indicating that rapamycin treatment hindered Ab secretion from long-lived plasma cells (58), our data suggest that in addition to Ig synthesis rates, the regulation of mTORC1 activity in long-lived plasma cells is governed at least in part by AMPK.

AMPK and mTORC1 reciprocally regulate autophagy, a conserved self-degrading cellular process (44). Autophagy is essential for the long-term persistence of both memory B and CD4 T cell populations (11, 12, 60). Loss of either two autophagy-essential genes, Atg5 or Atg7, was tied to mitochondrial dysfunction and enhanced lipid peroxidation in the B and T cell lineages (11, 60, 61). In a manner seeming analogous to our findings with AMPKα1, Atg5 in B cells was critical for limiting excessive Ig production by plasma cells (13), which was attributed to a function in restraining endoplasmic reticulum stress signaling. It was surprising, therefore, to observe that autophagy after glucose withdrawal was normal for AMPK-deficient B cells despite the expected absence of activating phosphorylation of ULK1. These findings suggest that activated B cells can also induce autophagy via noncanonical, AMPK-independent pathways (42). Despite normal formation of LC3 puncta, the defect in ULK1 (S317) phosphorylation was associated with a defect in ULK1 recruitment to the mitochondria and failure to induce mitophagy in AMPK-deficient B cells. In support of our findings, the AMPK–ULK1 axis is reported to be essential for hypoxia- or exercise-induced mitophagy in mouse embryonic fibroblasts and skeletal muscle cells, respectively (46, 47). Interestingly, previous studies demonstrating the importance of autophagy in lymphocyte persistence and mitochondrial homeostasis use genetic models by conditional inactivation of Atg5 or Atg7, both of which may be involved in multiple autophagy pathways, including mitophagy (42). Accordingly, the reason memory B cell persistence is promoted by Atg7 may be attributable in part to mitochondrial-specific autophagy activated via AMPKα1.

Our data show that AMPK promotes both the long-term persistence of memory B lymphocytes and mitophagy in B lineage cells. There are substantial antecedents that connect mitochondria to memory B cell persistence (62). Memory B cell longevity is linked to enhanced expression of mitochondrial prosurvival proteins in the Bcl2 superfamily, both Bcl2 itself (63) and BH3-only Puma (64). Furthermore, mitochondrial homeostasis is associated with the persistence of memory B cells (11, 62). Mitophagy has also been associated with maintaining memory in an NK cell population (65). In that setting, it likely functioned through clearance of dysfunctional mitochondria, which protected cells from the accumulation of excessive mtROS, lipid peroxidation, and cell death. Our findings with AMPK-deficient B cells align with this previous work in that we observed increased mitochondrial reactive oxygen species and abnormal function in mitochondrial testing during metabolic flux analyses. Spare oxidative phosphorylation, or the reserved capacity to amplify respiration in response to increased demand, was impaired in B cells lacking AMPK, similar to observations on memory CD8 T cell longevity (66). The types of memory lymphocytes assayed in prior work and ours are known to circulate through blood, tissue, and lymphatics (67). Accordingly, we speculate that spare respiratory capacity may be critical for memory cells to adapt to substantial differences in these microenvironments and the attendant metabolic stresses as they survey and pass through distinct tissues.

One observation in this work is that memory B cells and long-lived plasma cells exhibited distinct metabolic profiles and dependence on AMPK. Although AMPK supported the persistence of the memory B cell population, it dampened long-lived plasma cell Ab synthesis function without hindering survival. Activation of the AMPK pathway by metformin has been associated with increased memory B cells and improved Ab responses to influenza vaccine in type II diabetic patients (68). Our data may provide insight on the efficacy of drugs that target AMPK to achieve longer lasting humoral responses and improve vaccine design.

We thank Ariel Raybuck for sharing mice, techniques, and ideas throughout the duration of the work as well as careful readings of the manuscript. We thank Majan Rafat for assistance in quantification of confocal images, Kevin Chen for performing some ELISAs, Lan Wu for managing and troubleshooting FACSCanto equipment, and Linda Sealy and Janice Blum for guidance and support. We also thank the High-Throughput Screening, Cell Imaging Shared Resource, Flow Cytometry Shared Resource, and Cell and Developmental Biology Cores at Vanderbilt University and Vanderbilt University Medical Center for equipment, expertise, and assistance.

This work was supported by National Institutes of Health (NIH) Grants R01 AI113292 and R01 HL106812 (to M.R.B.). S.K.B. was supported by NIH Grants R25 GM062459 and T32 CA009592-29, followed by a supplement to R01 AI113292 and PMI Departmental funds. Additional support for S.K.B. was provided by Vanderbilt University’s Provost Graduate Fellowship Award. Additional support for P.J.B. was provided by Fundação de Amparo à Pesquisa do Estado de São Paulo 2018/08563-8. NIH Shared Instrumentation Grant 1S10OD018015 as well as scholarships via Cancer Center Support Grant CA068485 and Diabetes Research Center Grant DK0205930 helped defray costs of Vanderbilt Cores.

The online version of this article contains supplemental material.

Abbreviations used in this article:

AMPK

adenosine monophosphate–activated protein kinase

ASC

Ab-secreting cell

GC

germinal center

huCD20

human CD20

MphenBC

memory-phenotype B cell

mTORC1

mechanistic target of rapamycin complex 1

mtROS

mitochondrially derived reactive oxygen species

NP

nitrophenol

NP-KLH

4-hydroxy-3-nitrophenylacetyl hapten conjugated to keyhole limpet hemocyanin

OCR

oxygen consumption rate

4-OHT

4-hydroxytamoxifen

ULK1

Unc-51–like kinase.

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The senior author (M.R.B.) and coauthor S.H.C. received support, unrelated to the work described in this article, derived from a contract with Ribon Therapeutics, Inc. (Cambridge, MA), to test how the absence of PARP14 in our knockout mice affected growth, immune infiltration, or anti-PD1 responsiveness of orthotopic, triple-negative breast cancer cells in mice. In addition, the senior author (M.R.B.) holds various stocks ($5000–$20,000, depending on the day; FedEx, Amazon, Regeneron, etc.) in his own name through an investment advisor as part of overall savings for retirement, but none are in any identifiable conflict with this work. The senior author has accepted an offer to serve as expert witness for the defendant in a vaccine-autism tort. The other authors have no financial conflicts of interest.

Supplementary data