Article

Mast Cell-Derived Histami

ne Regulates LiverKetogenesis via Oleoylethanolamide Signaling

Graphical Abstract

Highlights

d Fasting stimulates extra-hepatic mast cells to release

histamine

d Histamine travels through the hepatic portal system and

activates liver H1 receptors

d H1-receptor activation triggers OEA biosynthesis in liver

d Histamine-dependent OEA signaling enhances fasting-

induced ketogenesis

Misto et al., 2019, Cell Metabolism 29, 91–102January 8, 2019 ª 2018 Elsevier Inc.https://doi.org/10.1016/j.cmet.2018.09.014

Authors

Alessandra Misto, Gustavo Provensi,

Valentina Vozella,

Maria Beatrice Passani,

Daniele Piomelli

Correspondencepiomelli@uci.edu

In Brief

Misto et al. describe a paracrine signaling

mechanism that enhances ketogenesis,

a crucial adaptive response to food

deprivation. The mechanism involves

fasting-induced release of histamine from

mast cells into the hepatic portal system,

stimulation of H1 receptor-dependent

OEA biosynthesis in liver, and

co-activation of PPAR-a-mediated

ketogenesis by OEA and lipolysis-derived

fatty acids.

Cell Metabolism

Article

Mast Cell-Derived HistamineRegulates Liver Ketogenesisvia Oleoylethanolamide SignalingAlessandra Misto,1,2 Gustavo Provensi,3 Valentina Vozella,1 Maria Beatrice Passani,4 and Daniele Piomelli5,6,*1Drug Discovery and Development, Fondazione Istituto Italiano di Tecnologia, Genoa 16163, Italy2School of Advanced Studies Sant’Anna, Pisa 56127, Italy3Department of Neuroscience, Psychology, Drug Research and Child Health, University of Florence, Florence 50139, Italy4Department of Health Science, University of Florence, Florence 50139, Italy5Departments of Anatomy and Neurobiology, Biological Chemistry and Pharmacology, School of Medicine, University of California, Irvine,

CA 92697, USA6Lead Contact*Correspondence: piomelli@uci.edu

https://doi.org/10.1016/j.cmet.2018.09.014

SUMMARY

The conversion of lipolysis-derived fatty acids intoketone bodies (ketogenesis) is a crucial metabolicadaptation to prolonged periods of food scarcity.The process occurs primarily in liver mitochondriaand is initiated by fatty-acid-mediated stimulationof the ligand-operated transcription factor, peroxi-some proliferator-activated receptor-a (PPAR-a).Here, we present evidence that mast cells contributeto the control of fasting-induced ketogenesis viaa paracrine mechanism that involves secretion ofhistamine into the hepatic portal circulation, stimula-tion of liver H1 receptors, and local biosynthesis ofthe high-affinity PPAR-a agonist, oleoylethanola-mide (OEA). Genetic or pharmacological interven-tions that disable any one of these events, includingmast cell elimination, deletion of histamine- or OEA-synthesizing enzymes, and H1 blockade, blunt keto-genesis without affecting lipolysis. The results revealan unexpected role for mast cells in the regulationof systemic fatty-acid homeostasis, and suggestthat OEA may act in concert with lipolysis-derivedfatty acids to activate liver PPAR-a and promoteketogenesis.

INTRODUCTION

After a meal, the absorptive epithelium of the upper small intes-

tine diverts a fraction of the oleic acid derived from the hydrolysis

of dietary lipids toward the production of oleoylethanolamide

(OEA) (Schwartz et al., 2008; DiPatrizio and Piomelli, 2015).

Acting as a local messenger within the gut, OEA reduces food

intake through a mechanism that involves recruitment of periph-

eral sensory afferents (Rodriguez de Fonseca et al., 2001) and

activation of central pathways that utilize oxytocin and histamine

as neurotransmitters (Gaetani et al., 2010; Provensi et al., 2014).

Several lines of evidence suggest that OEA initiates this

Ce

response by engaging the ligand-operated transcription factor,

peroxisome proliferator-activated receptor-a (PPAR-a). First,

OEA is one of the most potent naturally occurring PPAR-a

agonists identified to date (affinity constant, KD, z40 nM;

median effective concentration, EC50, z120 nM) (Fu et al.,

2003; Astarita et al., 2006a). Second, the satiety-inducing effects

of OEA are abolished by PPAR-a deletion, are mimicked by

synthetic PPAR-a agonists, and are associated with increased

PPAR-a-regulated transcription in gut mucosa (Fu et al., 2003).

Third, viral-mediated enhancement of OEA production in rat

jejunum reduces food intake and concomitantly increases local

expression of PPAR-a target genes (Fu et al., 2008). Finally,

OEA levels in small intestine from various vertebrate species,

including fish, snakes, and rodents, rise from %50 nM in the

fasting state to R250 nM after feeding (Astarita et al., 2006b;

Fu et al., 2007; Tinoco et al., 2014; Igarashi et al., 2017), a con-

centration range that is compatible with PPAR-a activation

(Fu et al., 2003; Astarita et al., 2006a). In sum, the available

data indicate that OEA is a physiologically relevant endogenous

agonist for small-intestinal PPAR-a, which participates in the

control of satiety (DiPatrizio and Piomelli, 2015).

Feeding regulates OEA production also in the liver, but in an

opposite direction to that seen in the gut: hepatic OEA levels

rise in the fasting state and fall after feeding (Fu et al., 2007;

Izzo et al., 2010). The molecular underpinnings and physiological

significance of these changes are unknown, but pharmacolog-

ical evidence hints at a possible role in lipid metabolism. Indeed,

subchronic administration of OEA attenuates liver steatosis in

obese Zucker rats (Fu et al., 2005) and in a rat model of non-

alcoholic fatty liver disease (Li et al., 2015). Moreover, exoge-

nous OEA stimulates fatty-acid oxidation in isolated rat

hepatocytes and enhances ketone body production (ketogen-

esis) in live rats (Guzman et al., 2004). Ketogenesis is an adaptive

metabolic response to prolonged nutrient insufficiency, which

takes place primarily in liver mitochondria (Grabacka et al.,

2016). The process starts with the condensation, catalyzed

by Acetyl-CoA acetyltranferase-1 (Acat-1), of two molecules

of acetyl-CoA to form acetoacetyl-CoA. Addition of another

acetyl-CoA group, mediated by b-Hydroxy-b-methylglutaryl-

CoA synthase-2 (Hmgcs-2), yields HMG-CoA, which is

then converted into the ketone bodies, acetoacetate and

ll Metabolism 29, 91–102, January 8, 2019 ª 2018 Elsevier Inc. 91

b-hydroxybutyrate (b-OHB). The expression of genes involved in

this pathway, including the rate-limiting enzyme Hmgcs-2

(Grabacka et al., 2016), is governed at the transcriptional level

by PPAR-a in hepatocytes (Montagner et al., 2016), whose acti-

vation in fasting animals is thought to depend primarily on lipol-

ysis-derived fatty acids (Kersten, 2014; Dubois et al., 2017).

Here, we describe a paracrine signaling mechanism involved in

the control of liver ketogenesis. We show that fasting stimulates

extra-hepatic mast cells to secrete histamine, which enters the

liver via the portal circulation, activates G protein-coupled H1

receptors, and triggers local biosynthesis of the high-affinity

PPAR-a agonist, OEA. Genetic or pharmacological manipula-

tions that disrupt this process do not affect lipolysis, but mark-

edly attenuate fasting-induced ketogenesis, thus revealing a

previously unsuspected regulatory role for mast cell-derived his-

tamine and liver OEA production in systemic lipid homeostasis.

RESULTS

Endogenous Histamine Drives Fasting-Induced OEABiosynthesis in LiverHistamine regulates food intake (Haas et al., 2008) andmediates,

along with oxytocin, the satiety response evoked by OEA (Gae-

tani et al., 2010; Provensi et al., 2016). In addition to these central

actions, histamine can also stimulate carbohydrate and lipid

metabolism by acting at peripheral sites (Garcia-Sainz et al.,

1987; Masaki et al., 2001a; Fulop et al., 2003; Wang et al.,

2010; Yamada et al., 2016). To assess whether OEA participates

in the peripheral metabolic effects of histamine, we examined

the impact of feeding status on OEA levels in mice lacking the

histamine-synthesizing enzyme, Histidine decarboxylase (Hdc)

(Ohtsu et al., 2001). Hdc�/� mice and their wild-type littermates

were subjected to three feeding conditions that are known to

influence OEA production in a tissue-selective manner (Fu

et al., 2007): free feeding, food deprivation (12 hr), and food

deprivation followed by 1 hr refeeding. Using liquid chromatog-

raphy/mass spectrometry (LC/MS), we measured OEA content

in four tissues: liver, white fat and pancreas (where fasting

elevates OEA levels), and jejunum (where the opposite occurs)

(Rodriguez de Fonseca et al., 2001; Fu et al., 2007; Schwartz

et al., 2008). Hdc ablation suppressed fasting-induced OEA

accumulation in liver (Figure 1A), but did not alter the OEA

response in jejunum (Figure 1B), fat (Figure 1C), or pancreas (Fig-

ure 1D). Moreover, Hdc deletion had no effect on hepatic levels

of the OEA analog palmitoylethanolamide (PEA) (Figure 1E) or on

lipolysis-mediated release of oleic acid (Figure 1F) and other free

fatty acids (FFAs) (Figure S1).

There are two known pathways of OEA biosynthesis (Piomelli,

2013). In gut mucosa, dietary oleic acid is condensed with

phosphatidylethanolamine (PE) to form N-oleoyl-PE, which is

converted to OEA by N-acyl-PE phospholipase D (Nape-pld)

(Fu et al., 2007, 2008; Schwartz et al., 2008; Piomelli, 2013).

Alternatively, in neurons and other cells, N-oleoyl-PE is gener-

ated via transfer of oleic acid from the sn-1 position of phospha-

tidylcholine to the free amine group of PE (Piomelli, 2013). This

reaction is initiated by activity- or receptor-dependent mobiliza-

tion of intracellular calcium (Cadas et al., 1996, 1997) and is

catalyzed by the N-acyltransferase Pla2g4e (Ogura et al.,

2016). To determine whether Hdc ablation impacts OEA biosyn-

92 Cell Metabolism 29, 91–102, January 8, 2019

thesis, we identified and quantified by LC/MS (Table S1)

three OEA precursors whose levels changed in response to

food deprivation: N-oleoyl-PE (18:0-22:6-N18:1), N-oleoyl-PE

(18:1-22:6-N18:1) and N-oleoyl-PE (16:0-18:1-N18:1). Fasting

caused aR3-fold increase in the accumulation of each of these

lipids, and Hdc deletion either suppressed (Figure 1G) or sub-

stantially dampened (Figures 1H and S2A) this response. By

contrast, neither fasting nor Hdc ablation had any effect on the

PEA precursors, N-palmitoyl-PE (18:0-22:6-N16:0) (Figure 1I)

and plasmalogen N-palmitoyl-PE (P18:0-22:6-N16:0) (Fig-

ure S2B). The results show that Hdc is required for fasting-

induced OEA biosynthesis in liver, implicating histamine as an

obligatory mediator of this process.

Histamine Stimulates Liver OEA Biosynthesis via H1

Receptor ActivationTo further test this idea, we examined whether pharmacological

blockade of histamine receptors affects liver OEA formation. His-

tamine activates with submicromolar potencies four G protein-

coupled receptors, termed H1 to H4 (Bongers et al., 2010).

Similar to Hdc ablation, injection of a maximally effective dose

of the H1 antagonist fexofenadine (10 mg/kg) (Matsubara et al.,

2017) prevented the fasting-induced rise in liver OEA levels

without changing such levels in fedmice (Figure 2A). By contrast,

the H2 antagonist famotidine (10mg/kg) (Swiader and Czuczwar,

2014) and the H3 antagonist ciproxifan (3 mg/kg) (Huang et al.,

2006) had no effect on OEA production under any feeding condi-

tion (Figures 2B and 2C), while the H4 antagonist JNJ7777120

(20 mg/kg) (Ballerini et al., 2013) slightly raised OEA content in

free-feeding, but not food-deprived mice (Figure 2D). Further-

more, supporting the possibility that histamine regulates OEA

levels via a liver-autonomousmechanism, we found that (1) intra-

hepatic histamine injections increased liver OEA content in fed

animals (Figure 2E), and (2) incubation in the presence of hista-

mine stimulated OEA formation in the human hepatocyte cell

line, HepG2 (Figure 2F), which expresses H1 receptors (Falus

et al., 1989). Conversely, intracerebroventricular infusion of the

irreversible Hdc inhibitor, a-fluoromethylhistidine (a-FMH) (Gar-

barg et al., 1980), which effectively depletes releasable hista-

mine stores in the mouse brain (Provensi et al., 2014), did not

change fasting-induced OEA production (Figure 2G). The evi-

dence thus suggests that endogenous histamine stimulates

OEA biosynthesis in liver via H1 receptor activation.

Fasting Stimulates Histamine Release from Mast Cellsinto the Hepatic Portal SystemNext, we asked whether fasting changes histamine levels in the

hepatic portal circulation, which provides a likely point of entry

for the amine into the liver. The results show that mouse portal

blood contains measurable levels of histamine and that fasting

markedly increased such levels (Figures 3A and 3B). As ex-

pected from previous reports (Ohtsu et al., 2001; Nakamura

et al., 2017), histamine was also detectable in portal blood of

Hdc�/� mice, most likely due to dietary intake (Ohtsu et al.,

2001, 2002). In these mutants, however, histamine levels did

not change in response to food deprivation (Figure 3C).

Outside the brain, histamine is found predominantly in mast

cells, basophils, and gastric enterochromaffin-like cells, from

which it can be released in response to a variety of immune,

Figure 1. Endogenous Histamine Is Required for Fasting-Induced OEA Biosynthesis in Liver

Effects of Hdc deletion on levels of OEA in (A) liver, (B) jejunum, (C) epididymal white adipose tissue (WAT), and (D) pancreas. Hdc deletion does not affect liver

levels of (E) PEA or (F) oleic acid. Biosynthesis of OEA precursors (G) N-oleoyl-PE (18:0-22:6-N18:1) and (H) N-oleoyl-PE (18:1-22:6-N18:1) is impaired in Hdc�/�

mice, whereas biosynthesis of the PEA precursor, (I) N-palmitoyl-PE (18:0-22:6-N16:0), is unaltered. Wild-type (WT) and Hdc�/� mice were tested under three

experimental conditions: free feeding (open bars), food deprivation (12 hr, gray bars), and food deprivation followed by 1 hr refeeding (closed bars). Results are

expressed as mean ± SEM, n = 5–6 per feeding condition.

*p < 0.05, **p < 0.01, ***p < 0.001. See also Figures S1 and S2; Table S1.

hormonal, and neural signals (Prinz et al., 1999; Borriello et al.,

2017). Because a substantial fraction of blood draining into the

portal vein derives from the gastrointestinal tract, which harbors

a functionally distinct population of mast cells (Pearce, 1986;

Stead et al., 2006), we asked whether histamine release in portal

blood and liver OEA biosynthesis might be altered in mice lack-

ing these cells. In a model of Kit-dependent mast cell deficiency,

WBB6F1/J-KitW/KitW-v/Jmice (Grimbaldeston et al., 2005), food

deprivation failed to affect portal histamine concentrations (Fig-

ure 4A) or liver OEA biosynthesis (Figures 4B and 4C). The deficit

was biochemically selective in thatKitmutants did not differ from

wild-type littermates in other responses evoked by fasting,

including lowered OEA production in jejunum (Figure 4D),

enhanced OEA production in adipose tissue (Figure 4E), and

accelerated lipolysis (Figure 4F). Because abnormalities in Kit

mutant mice extend beyondmast cell deficiency, encompassing

a wide variety of immunological and metabolic functions (Feyer-

abend et al., 2012, 2016), we also used Carboxypeptidase A3

(Cpa3) Cre recombinase knock-in (Cpa3Cre/+) mice, in which

Cre-mediated genotoxicity causes complete mast cell elimina-

tion along with anz60% reduction in basophil number (Feyera-

bend et al., 2011). As observed inKit-dependentmutants, fasting

Cell Metabolism 29, 91–102, January 8, 2019 93

(legend on next page)

94 Cell Metabolism 29, 91–102, January 8, 2019

Figure 3. Fasting Stimulates Histamine Release into the Hepatic Portal Circulation

Histamine identification and quantification in hepatic portal blood.

(A) Representative LC/MS tracing of authentic histamine (red) and native histamine (black).

(B) Mass spectrum of native histamine showing the presence of diagnostic ions [M + H]+ and [M-NH3]+.

(C) Effects of free feeding (open bars), food deprivation (gray bars), and food deprivation/refeeding (closed bars) on histamine levels in portal blood of wild-type

(WT) and Hdc�/� mice. Results are expressed as mean ± SEM, n = 4–5 per feeding condition. ***p < 0.001; #p < 0.01 versus corresponding group of WT mice.

caused no detectable change in histamine or OEAmobilization in

Cpa3Cre/+ mice (Figures 4G and 4H). Collectively, we interpret

these findings as indicating that food deprivation causes

extra-hepatic mast cells to secrete histamine, which reaches

the liver via the portal system and stimulates OEA biosynthesis

in that organ.

Dietary Fat Dampens Histamine Release and Liver OEAProductionIn the small intestine, where OEA acts as a satiety factor

(DiPatrizio and Piomelli, 2015), the feeding-dependent formation

of this lipid mediator is disabled by short-term or long-term

exposure to a high-fat diet (HFD) (Artmann et al., 2008; Igarashi

et al., 2015). To determine whether dietary fat also influences

histamine-dependent OEA biosynthesis in liver, we placed

wild-type mice on an HFD for 7 days and measured portal

histamine and hepatic OEA content under free-feeding, food-

deprived, and refed conditions. The short HFD exposure did

not change body weight (day 0: 25.9 ± 0.4 g; day 7: 27.0 ±

0.5 g; mean ± SEM; n = 15), but dampened fasting-induced

histamine release into the portal circulation (Figure S3A) and

suppressed OEA production in liver (Figure S3B). This effect

was temporary, however, as both fasting-induced responses

were normalized after 30 days of HFD (Figures S3C and

S3D), when mice exposed to the diet were significantly

(***p < 0.0001) heavier than chow-fed controls (standard diet,

day 30: 27.47 ± 0.37 g; HFD, day 30: 32.73 ± 0.63 g; mean ±

SEM; n = 15). Thus, histamine-dependent OEA formation in liver

is subject only to short-term but not long-term modulation by

Figure 2. H1 Receptors Mediate Fasting-Induced OEA Biosynthesis in

Effects of histamine receptor blockade on liver OEA levels in free-feeding (ope

(closed bars).

(A–F) (A) H1 antagonist fexofenadine (10 mg/kg), (B) H2 antagonist famotidine

JNJ7777120 (20mg/kg). Drugs were administered by intraperitoneal injection 1 hr

liver OEA levels in free-feeding wild-type mice, and (F) incubation of HepG2 cells

(G) Effect of intracerebroventricular infusion of the irreversible Hdc inhibitor, a

expressed as mean ± SEM, n = 5–6 per condition.

*p < 0.05, **p < 0.01, ***p < 0.001; #p < 0.01, free-feeding vehicle-treatedmice vers

versus vehicle.

dietary fat, suggesting that adaptive mechanisms might exist

that restore the response after prolonged fat feeding.

Histamine-Driven OEA Biosynthesis Enhances Fasting-Induced KetogenesisExperiments in rats have shown that exogenous OEA promotes

fatty-acid oxidation and ketone body formation by activating

PPAR-a (Guzman et al., 2004). These data prompted us to inves-

tigate the impact of Hdc deletion and histamine receptor antago-

nism on fasting-induced ketogenesis. In wild-type mice, food

deprivation increased b-OHB levels in liver and plasma (Figures

5A and 5B) and concomitantly stimulated transcription of the

ketogenic genes, Acat-1 and Hmgcs-2 (Figures 5C and 5D).

Expression of two additional PPAR-a targets, Fibroblast growth

factor 21 (Fgf21) and Cytochrome P450-4a10 (Cyp4a10), was

also enhanced (Figures 5E and 5F). Consistent with a regulatory

role for histamine, these responses were significantly attenuated

in Hdc�/� mice (Figures 5A–5F). Of note, Hdc deletion differen-

tially affected Fgf21 and Cyp4a10 transcription (Figures 5E and

5F). Similar discrepancies have been documented previously

(e.g., between fasting- and agonist-induced expression of

PPAR-a target genes) and have tentatively attributed to factors

such as cellular PPAR-a localization, concomitant engagement

of other nuclear receptors, and/or differential recruitment of

coactivator proteins (Kersten, 2014). As seen in Hdc�/� mice,

fasting-induced changes in hepatic and circulating b-OHB levels

were also blunted in wild-typemice treatedwith the H1 antagonist

fexofenadine (Figures 5G and 5H), but not in mice treated with

H2, H3, or H4 antagonists (Figures S4A–S4C). In addition,

Liver

n bars), food-deprived (gray bars), and food-deprived/refed wild-type mice

(10 mg/kg), (C) H3 antagonist ciproxifan (3 mg/kg), and (D) H4 antagonist

before tissue harvest. Effects of (E) intrahepatic injection of histamine (mmol) on

with histamine (30 mM, 1 hr).

-FMH, on OEA formation in liver of free-feeding wild-type mice. Results are

us free-feeding drug-treatedmice, except (E and F) where **p < 0.01, histamine

Cell Metabolism 29, 91–102, January 8, 2019 95

Figure 4. Mast Cells Are a Primary Source of Fasting-Induced Histamine Release

Effects of Kit-dependent (WBB6F1/J-KitW/KitW-v/J) or Kit-independent (Cpa3Cre/+) mast cell ablation on fasting-induced histamine release and liver OEA

biosynthesis.WBB6F1/J-KitW/KitW-v/Jmice: (A) histamine concentrations in portal blood; liver levels of (B) OEA and (C) N-oleoyl-PE (18:0-22:6-N18:1); levels of

OEA in (D) jejunum and (E) white adipose tissue (WAT); (F) liver levels of oleic acid. Cpa3Cre/+ mice: (G) histamine concentrations in portal blood; (H) liver OEA

levels. Free feeding (open bars), food deprivation (gray bars), food deprivation/refeeding (closed bars). Results are expressed asmean ± SEM, n = 4–5 per feeding

condition.

*p < 0.05, **p < 0.01, ***p < 0.001 versus wild-type (WT) littermate controls.

fexofenadine administration reduced fasting-inducedAcat-1 and,

to a lesser extent, Hmgcs-2 transcription (Figures S5A and S5B).

As illustrated (Figures 3 and 4), fasting failed to increase portal

histamine and liver OEA levels inKit-independent (Cpa3Cre/+) and

Kit-dependent (WBB6F1/J-KitW/KitW-v/J) models of mast cell

ablation. In the same strains, ketogenesis was also defective

(Figures 6A and 6B). Liver transcription of ketogenic (Acat-1

and Hmgcs-2) and other PPAR-a target genes (Fgf21 and

Cyp4a10) was similarly, albeit differentially, blunted (Figures

6C–6F and S6). If OEA is involved in ketone body production,

as the results outlined previously suggest, the process should

96 Cell Metabolism 29, 91–102, January 8, 2019

be impaired in mice lacking Nape-pld, the enzyme that converts

N-oleoyl-PE into OEA. Confirming this expectation, fasting-

induced rises in b-OHB levels were significantly decreased in

Nape-pld�/� mutants, compared with wild-type littermates (Fig-

ures 6G and 6H). Of note, Nape-pld ablation did not modify the

ketogenic response evoked by injections of OEA (20 mg/kg),

which increased liver b-OHB content to a comparable extent in

Nape-pld�/� andwild-type animals (Figure 6I). In sum, the results

outlined previously suggest that mast cell-derived histamine

magnifies fasting-induced ketogenesis by activating H1

receptor-dependent OEA production in liver.

Figure 5. Endogenous Histamine Enhances Fasting-Induced KetogenesisEffects ofHdc deletion and H1 receptor blockade on b-OHB levels in (A) liver and (B) plasma ofHdc�/�mice and wild-type (WT) controls under free-feeding (open

bars), food-deprived (gray bars), and food-deprived/refed (closed bars) conditions. Transcription of PPAR-a target genes (C) Acat-1, (D)Hmgcs-2, (E) Fgf21, and

(F) Cyp4a10 in liver tissue of WT and Hdc�/� mice. Effects of H1 antagonist fexofenadine (10 mg/kg, intraperitoneal) or vehicle on b-OHB levels in (G) liver and (H)

plasma of WT mice. Fexofenadine was administered 1 hr before tissue harvest. Results are expressed as mean ± SEM, n = 5 per feeding condition.

*p < 0.05, **p < 0.01, ***p < 0.001. See also Figures S4 and S5.

DISCUSSION

The present findings outline a novel regulatory mechanism for

fasting-induced ketogenesis, which involves histamine release

frommast cells into the hepatic portal system, H1 receptor-medi-

ated stimulation of liver OEA biosynthesis, and recruitment of the

nuclear receptor PPAR-a by the combined action of OEA and

lipolysis-derived FFAs (Figure 7). Three sets of data support

this model. First, fasting increases portal histamine levels in

wild-type mice, but not in three mutant mouse strains that lack

either mast cells or Hdc. Second, fasting is accompanied by a

marked enhancement of liver OEA production, a response that

is suppressed by mast cell removal, genetic Hdc silencing, or

pharmacological H1 receptor blockade. Finally, experimental

interventions that disable histamine release and/or liver OEA

biosynthesis also attenuate, but do not completely block, fast-

ing-induced PPAR-a activation and ketogenesis. Notably, the

same interventions have no effect on the accumulation of lipol-

ysis-derived FFAs.

We found that fasting increases histamine levels in portal

blood and concomitantly heightens liver OEA production. These

responses are suppressed in two mechanistically distinct

models of mast cell deficiency: Cpa3Cre/+ mice, in which mast

cells are eliminated by Cre genotoxicity (Feyerabend et al.,

2011), and WBB6F1/J-KitW/KitW-v/J mice, in which mast cell

ablation is caused by impaired Kit signaling and is accompanied

by various immune and metabolic deficits (Gutierrez et al., 2015;

Feyerabend et al., 2016). Collectively, the results support the

hypothesis that mast cells are critically implicated in fasting-

induced histamine secretion. Of note, portal histamine levels

undergo an z200% change in the transition from feeding to

fasting (Figures 4A and 4G). This increase is comparable in

size to those observed in peripheral plasma of asthmatic patients

experiencing challenge-induced bronchoconstriction (Bhat

et al., 1976; Belcher et al., 1988), an event in which histamine

is known to play an important functional role (Kaliner, 1989).

The specific mast cell population(s) involved in fasting-induced

histamine release remains to be identified, but the available evi-

dence points to three candidate sites: gastrointestinal mucosa,

visceral vasculature, and/or portal tracts and sinusoids of the

liver (Farrell et al., 1995; Stead et al., 2006). The distinctive pop-

ulation of mast cells that resides in the gut may be well posi-

tioned to fill this role. Indeed, mucosal mast cells are found in

close proximity of neural fibers (Stead et al., 1987, 1989),

including vagal efferents that are activated during a fast

(Berthoud and Patterson, 1996; Berthoud, 2008a, 2008b). More-

over, mucosal mast cells respond to vagal stimulation with

increased histamine biosynthesis (Stead et al., 2006) and have

been implicated in the monitoring and utilization of dietary fat

(Ji et al., 2012; Sato et al., 2016). Further investigations are

needed to test these possibilities.

The stimulus-dependent production of OEA, like that of most

lipid-derived mediators, is regulated in a cell- and tissue-

selective manner (Piomelli, 2013). Unlike gut mucosa, where

OEA is produced using dietary oleic acid as an obligatory precur-

sor (Schwartz et al., 2008), in the liver this process is ostensibly

uncoupled from influx of lipolysis-derived FFAs (compare

Figures 1A and 1F). It is possible, therefore, though remains to

be demonstrated, that liver cells produce OEA through a

pathway similar to that used by brain neurons, namely, via

Cell Metabolism 29, 91–102, January 8, 2019 97

Figure 6. Role of Mast Cells and OEA in Fasting-Induced Ketogenesis

(A–H) Effects of Kit-independent (Cpa3Cre/+) or Kit-dependent (WBB6F1/J-KitW/KitW-v/J) mast cell ablation on plasma b-OHB levels in (A) Cpa3Cre/+ mice and (B)

WBB6F1/J-KitW/KitW-v/J mice under free-feeding (open bars), food-deprived (gray bars), and food-deprived/refed (closed bars) conditions. Transcription of

PPAR-a target genes (C)Acat-1, (D)Hmgcs-2, (E) Fgf21, and (F)Cyp4a10 in liver ofCpa3Cre/+mice. Effects ofNape-pld deletion on (G) liver and (H) plasma b-OHB

levels.

(I) Effects of intraperitoneal injections of OEA (20mg/kg, green bars) or vehicle (open bars) on b-OHB levels in liver of free-feeding wild-type (WT) andNape-pld�/�

mice. OEA was administered 2 hr before tissue harvest. Results are expressed as mean ± SEM, n = 5 per feeding condition.

*p < 0.05, **p < 0.01, ***p < 0.001 versus WT littermate controls. See also Figure S6.

mobilization of intracellular calcium and subsequent activation of

Pla2g4e, a phospholipase/N-acyltransferase responsible for

N-oleoyl-PE production (Piomelli, 2013; Ogura et al., 2016).

Consistent with this view, previous studies have documented

the presence of functionally active H1 receptors linked to calcium

signaling in hepatocytes (Garcia-Sainz et al., 1992) and other

cellular constituents of the liver (e.g., endothelial cells) (Naka-

mura andMurata, 2018). The identification of specific cell type(s)

involved in histamine-driven OEA formation will allow testing this

hypothesis and is an important objective for future studies.

98 Cell Metabolism 29, 91–102, January 8, 2019

Mammals adapt to prolonged periods of food scarcity by

releasing FFAs from adipose tissue and transforming them into

energy-rich ketone bodies that are used as fuel by brain, muscle,

and other organs (Kersten, 2014; Grabacka et al., 2016). Keto-

genesis occurs primarily in hepatocytes, but its rate is regulated

by a combination of local and systemic factors. Systemic sig-

nals, such as the catabolic hormones glucagon and epinephrine,

accelerate lipolysis and facilitate FFA transport into hepatocytes,

where these lipids may serve as both precursors for ketone

bodies and agonists for the master transcriptional regulator of

Figure 7. Mast Cell-Derived Histamine Reg-

ulates Ketogenesis via OEA Signaling

According to this model, fasting stimulates extra-

hepatic mast cells to secrete histamine into the

portal circulation. The amine enters the liver

where it triggers OEA biosynthesis by interacting

with Gq-coupled H1 receptors (H1R), which

activate the N-oleoyl-PE (NOPE)-synthesizing

enzyme Pla2g4e via intracellular calcium mobili-

zation. Nape-pld converts N-oleoyl-PE into the

high-affinity PPAR-a agonist, OEA. Newly formed

OEA cooperates with lipolysis-derived FFAs

(and/or other endogenous ligands) to activate

PPAR-a, the master transcriptional regulator of

ketogenesis. TAG, triacylglycerols.

ketogenesis, PPAR-a (Montagner et al., 2016; Dubois et al.,

2017). Nevertheless, the fact that FFAs activate this nuclear re-

ceptor only at mid to high micromolar concentrations (Gottlicher

et al., 1992; Berger andMoller, 2002), which are unlikely to occur

in healthy cells (Yamashita et al., 1997), challenges a simplistic

interpretation of this model and suggests that additional ligands

might be implicated in fasting-induced activation of PPAR-a in

liver (Chakravarti, 2009; Kersten, 2014). Our results show that

ketogenesis is substantially dampened, but not completely sup-

pressed, by genetic or pharmacological manipulations that

disrupt histamine-driven OEA biosynthesis. A plausible interpre-

tation of this finding is that OEA acts in concert with FFAs

(and/or, possibly, other endogenous ligands) (Chakravarti,

2009) to engage PPAR-a and promote ketogenesis (Figure 7).

Histamine-releasing neurons housed in the tuberomamillary

nucleus of the hypothalamus regulate circadian feeding and

mediate, in part, the satiety response initiated by OEA mobiliza-

tion in the small intestine (Provensi et al., 2014; DiPatrizio and

Piomelli, 2015). In addition to these central anorexic effects,

Cell M

which depend on H1 receptor activation

(Haas et al., 2008), histamine also contrib-

utes in important ways to the peripheral

control of energy balance and lipid meta-

bolism. Indeed, in mice, genetic H1 recep-

tor deletion causes increased abdominal

adiposity and glucose intolerance (Ma-

saki et al., 2001b), while pharmacological

H1 blockade aggravates HFD-induced

hepatic steatosis (Raveendran et al.,

2014). Similarly, in obese humans,

chronic use of H1 antagonists, which are

commonly prescribed to alleviate allergy

symptoms, is associated with increased

weight gain and insulin resistance (Ratliff

et al., 2010). H1 blockade has been also

implicated in the weight gain and dyslipi-

demia that often accompany the pro-

tracted use of atypical antipsychotic

drugs (Nasrallah, 2008). The mecha-

nism(s) underlying these dysmetabolic

actions is unclear. However, our finding

that the H1 antagonist fexofenadine

disables histamine-dependent signaling

and impairs ketogenesis points to a possible explanation, which

warrants, of course, further investigation.

Limitations of StudyThe present study has several limitations. First, while we demon-

strated that fasting stimulates histamine release frommast cells,

we did not identify the neuro-humoral signal(s) and specific mast

cell population(s) involved in this response. Second, we did not

define the liver cell type(s) involved in histamine-driven OEA

signaling and, consequently, we cannot draw firm conclusions

about the molecular mechanism through which H1 receptor

activation stimulates OEA production. Third, we showed that

histamine-dependent OEA production in liver contributes to

fasting-induced ketogenesis, but did not clarify how OEA

interacts with other endogenous PPAR-a ligands (e.g., FFAs)

to produce this effect. Finally, the potential contribution of

histamine-driven OEA signaling to metabolic disease was not

investigated. Despite these limitations, our experiments reveal

an unexpected new role for mast cell-derived histamine in the

etabolism 29, 91–102, January 8, 2019 99

regulation of fasting-induced liver ketogenesis, a key node of

bioenergetics homeostasis, and raise the intriguing possibility

that dysfunctions in this paracrine signaling mechanism might

contribute to metabolic disorders.

STAR+METHODS

Detailed methods are provided in the online version of this paper

and include the following:

d KEY RESOURCES TABLE

d CONTACT FOR REAGENT AND RESOURCE SHARING

d EXPERIMENTAL MODEL AND SUBJECT DETAILS

B Animals

B Cell Cultures

d METHOD DETAILS

B Experimental Design

B Tissue Collection and Plasma Preparation

B Drug Administrations

B Intrahepatic Histamine Injections

B Intracerebroventricular Infusion of a-FMH

B Lipid Extraction and Fractionation

B LC/MS Analyses

B Ketone Body Measurements

B Gene Expression Analyses

B Chemicals

d QUANTIFICATION AND STATISTICAL ANALYSIS

SUPPLEMENTAL INFORMATION

Supplemental Information includes six figures and one table and can be found

with this article online at https://doi.org/10.1016/j.cmet.2018.09.014.

ACKNOWLEDGMENTS

We thank Dr. T. Feyerabend and Prof. H.R. Rodewald for Cpa3Cre/+ mutant

mice and critical reading of the manuscript; Drs. S. Galli, S. Pontis, N. Realini,

F. Palese, and P. Blandina for advice and support; Drs. A. Basit and M.E.

Blanco for LC/MS analyses; and A. Costa for help with sample collection.

AUTHOR CONTRIBUTIONS

A.M. performed the experiments, analyzed the data, prepared figures and ta-

bles. G.P. and V.V. provided help with some animal experiments. M.B.P.

participated in experimental design. D.P. ideated and overviewed the project,

and wrote the manuscript with contributions from all coauthors. All authors

reviewed the results and approved the final version of the manuscript.

DECLARATION OF INTERESTS

The authors declare no competing interests.

Received: July 18, 2017

Revised: June 11, 2018

Accepted: September 12, 2018

Published: October 11, 2018

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cylases involved in fatty acid remodeling of phospholipids and metabolism of

bioactive lipids in mammalian cells. J. Biochem. 122, 1–16.

STAR+METHODS

KEY RESOURCES TABLE

REAGENT or RESOURCE SOURCE IDENTIFIER

Chemicals, Peptides, and Recombinant Proteins

1-stearoyl, 2-docosahexaenoyl-phosphatidylethanolamine

N-heptanoyl (N-heptanoyl-PE)

(Basit et al., 2016) N/A

Ciproxifan hydrochloride Sigma-Aldrich Cat#C6492; CAS: 1049741-81-2

Famotidine Sigma-Aldrich Cat#F6889; CAS: 76824-35-6

Fatty acids Sigma-Aldrich Cat#EC10A

Fexofenadine hydrochloride Sigma-Aldrich Cat#F9427; CAS: 153439-40-8

Heparin sodium salt from porcine intestinal mucosa Sigma-Aldrich Cat#H4784-1G; CAS: 9041-08-1

Heptadecanoic acid Sigma-Aldrich Cat#H3500; CAS: 506-12-7

Histamine Sigma-Aldrich Cat#H7125; CAS: 51-45-6

Histamine-a,a,b,b- d4 dihydrochloride ([2H4]-histamine) Sigma-Aldrich Cat#762962

JNJ7777120 Johnson & Johnson Reference Number: 1010888584

L-Glutamine 100X (200mM) EuroClone S.p.A. ECB3000D-20

Oleoyl Ethanolamide (OEA) Cayman Chemicals Cat#90265; CAS: 111-58-0

Oleoyl Ethanolamide-d4 ([2H4]-OEA) Cayman Chemicals Cat#9000552; CAS: 946524-36-3

Palmitoyl Ethanolamide (PEA) Cayman Chemicals Cat#90350; CAS:544-31-0

Palmitoyl Ethanolamide-d4 ([2H4]-PEA) Cayman Chemicals Cat#10007824; CAS: 1159908-45-8

Trypsin-EDTA Sigma-Aldrich T4049-500ML

a-FMH Johnson & Johnson Reference Number: 9101053137

Critical Commercial Assays

iTaqTM Universal SYBR Green Supermix Bio Rad Cat#172-5124 25ML

PureLinkTM DNase Thermo Fisher Scientific Cat#12185-010

PureLinkTM RNA Mini Kit Thermo Fisher Scientific Cat#12183-018A

SuperScript VILOTM cDNA Synthesis Kit Thermo Fisher Scientific Cat#11754-050

b-OHB (Ketone Body) Colorimetric Assay Kit Cayman Chemicals Cat#700190

Experimental Models: Cell Lines

Human hepatocyte cell line (HepG2) Sigma-Aldrich 85011430

Experimental Models: Organisms/Strains

Mouse: C57Bl/6J Charles River JAX 000664

Mouse: Cpa3Cre/+ German Cancer Research Center,

Heidelberg; (Feyerabend et al., 2011)

N/A

Mouse: Hdc-/- CeSAL, University of Florence;

(Ohtsu et al., 2001; Provensi et al., 2014)

N/A

Mouse: Nape-pld-/- University of California, Irvine;

(Leung et al., 2006)

N/A

Mouse: WBB6F1/J-KitW/KitW-v/J The Jackson Laboratory JAX 100410

Oligonucleotides

Mouse Acat-1 forward primer

5’-GTCTGGCTAGTATTTGCAACG-3’

This paper N/A

Mouse Acat-1 reverse primer

5’-TTCAGCCGGTCACATGG-3’

This paper N/A

Mouse Cyp4a10 forward primer

5’-GCTACTCAAGGCTTTCCAGCAG-3’

This paper N/A

Mouse Cyp4a10 reverse primer

5’-CCAGAACCATCTAGGAAAGGCAC-3’

This paper N/A

Mouse Fgf21 forward primer

5’-ATCAGGGAGGATGGAACAGTGG-3’

This paper N/A

(Continued on next page)

Cell Metabolism 29, 91–102.e1–e5, January 8, 2019 e1

Continued

REAGENT or RESOURCE SOURCE IDENTIFIER

Mouse Fgf21 reverse primer

5’-AGCTCCATCTGGCTGTTGGCAA-3’

This paper N/A

Mouse Hmgcs-2 forward primer

5’-TGGTTCAAGACAGGGCACACAGAAC-3’

This paper N/A

Mouse Hmgcs-2 reverse primer

5’-AGAGGAATACCAGGGCCCAACAAT-3’

This paper N/A

Software and Algorithms

GraphPad Prism version 5.03 GraphPad Software https://www.graphpad.com/scientific-

software/prism/

Other

Rodent Diet with 60 kcal% fat, 5.24 kcal/g Research Diets D12492

Standard chow diet, 2.66 kcal/g Mucedola S.R.L. 4RF21 GLP

Advanced Minimum Essential Medium (MEM) Thermo Fisher Scientific Cat#12492-013

Fetal Bovine Serum (FBS) Thermo Fisher Scientific Cat#10270-500ML

TRIzolTM Reagent Thermo Fisher Scientific Cat#15596-018

CONTACT FOR REAGENT AND RESOURCE SHARING

Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Daniele

Piomelli (piomelli@uci.edu).

EXPERIMENTAL MODEL AND SUBJECT DETAILS

AnimalsWe obtained adult male C57Bl/6J mice (8-9 weeks, 25-30 g) from Charles River (Lecco, Italy). Male Hdc-/- mice and their congenic

WT littermates [129/Sv background; see (Provensi et al., 2014) for genotyping procedures] were bred in the Centro Stabulazione

Animali di Laboratorio (CeSAL, University of Florence, Italy) and used at 8-12 weeks of age (25-35 g). Male mast cell-deficient

(WBB6F1/J-KitW/KitW-v/J) mice (7-8 weeks, 20-25 g) were purchased from Jackson Laboratories (Bar Harbor, USA). Male

Cre-mediated mast cell eradication (Cpa3Cre/+) mice and their congenic WT littermates (Feyerabend et al., 2011) were bred at the

German Cancer Research Center (Heidelberg, Germany) and used at 15-20 weeks of age (25-33 g). Male Nape-pld-/- mice and their

congenic WT littermates [129/Sv background; see (Leung et al., 2006) for generation of mice] were obtained from the University

of California, Irvine, and were bred at the Italian Institute of Technology (Genoa, Italy). They were used at 8-9 weeks of age

(20-25 g). All mice were housed in groups of 5 in ventilated cages with free access to food and water, unless indicated otherwise.

They were maintained under a 12 h light/dark cycle (lights on at 07:00 AM) at controlled temperature (22 ± 1�C) and relative humidity

(55 ± 10%), and handled for 1 week before experiments. Housing, animal maintenance and all procedures were conducted in

accordance with European Union (O.J. of E.C. L 358/1 12/18/1986) and Italian (D.M. 116192) regulations on the protection of animals

used for experimental and other scientific purposes.

Cell CulturesThe hepatocyte human cell line HepG2was purchased fromSigma-Aldrich (Milan, Italy) and cultured in AdvancedMinimumEssential

Medium (MEM) supplemented with 10% fetal bovine serum (FBS), 2 mM L-glutamine and antibiotics (100 U/mL penicillin and

streptomycin) at 37�C and 5% CO2. Cells were seeded in 100 mm Petri dishes (1.5 x 106 cell/dish) with 8 mL complete medium

(MEM plus 10% FBS) and allowed to adhere overnight. On the following day, the cells were incubated at 37�C in the presence of

histamine (30 mM) or vehicle (80% sterile saline). After one hour, the medium (8 mL) was collected and stored at -80�C for OEA

quantification (see below). All experiments were performed with at cell passages 5–7.

METHOD DETAILS

Experimental DesignMice were housed in individual bottom-wired cages and habituated to the experimental setting for 4 days in 3 randomly assigned

groups: a) free feeding, b) 12 h food deprivation (from 7 PM to 7 AM) and c) 1h refeeding after food deprivation. Water and standard

chow pellets (2.66 kcal/g, 4RF21; SettimoMilanese, Italy) were available ad libitum except when animals were food deprived. In some

experiments, mice were randomly divided into two groups – standard chow diet or HFD (60 kcal% fat, 5.24 kcal/g, D12492 Research

Diets, Inc.; Gentofte, Denmark) – whichwere then allowed to feed ad libitum for 7 or 30 days. At the end of the experiment, the animals

e2 Cell Metabolism 29, 91–102.e1–e5, January 8, 2019

were anesthetized with isoflurane (selected because of its minimal impact on liver metabolism) (Overmyer et al., 2015; Takahashi

et al., 1998) and killed by cervical dislocation.

Tissue Collection and Plasma PreparationWe collected blood from the portal vein of anesthetized mice using a 1 mL syringe with 22G needle, and immediately transferred it to

1.5 mL vials containing heparin (0.02 mL). The blood samples were centrifuged for 15 min at 1,000xg and 4�C, plasma layers were

collected and stored at -80�C for further processing. Liver, epididymal white adipose tissue, pancreas and small intestine (jejunum)

were removed immediately, snap-frozen in liquid nitrogen and stored at -80�C.

Drug AdministrationsHistamine antagonists and OEA were administered by intraperitoneal (i.p.) injection 1 h and 2 h, respectively, before sacrifice.

Histamine antagonists (except JNJ7777120) were administered in sterile saline, JNJ7777120 in saline containing 1% ethanol, and

OEA in 80% sterile saline, 10% PEG 400, and 10% Tween 80.

Intrahepatic Histamine InjectionsFor intrahepatic injection, free-feedingmicewere placed in a supine position with the nose andmouth positioned inside amouthpiece

that delivered isoflurane [2% (v/v) in 1 L/min O2]. After disinfection of the surgical area, the skin was removed and a midline incision

was made through the upper abdomen and peritoneum. A portion of the liver was exposed and histamine (0.45 mmol or 0.9 mmol in

sterile saline) was slowly injected into the liver parenchyma using an insulin syringe (injection volume 0.1 mL). 30 min later, the mice

were killed and their livers were collected.

Intracerebroventricular Infusion of a-FMHAnesthesia was performed using a mixture of ketamine (15 mg/kg) and xylazine (2.5 mg/kg). Anesthesized mice were placed in a

stereotaxic frame (Kopf Instruments, Tujunga, USA) and a stainless steel cannula (7 mm in length, outer diameter 0.5 mm, and inner

diameter 0.25 mm) was implanted into the lateral cerebral ventricle and fixed to the skull using dental cement. We used the following

coordinates: antero-posterior (AP) -0.3; lateral (L) ± 1; dorsoventral (DV) -1. A stainless steel microinjection needle was connected

through a polyethylene catheter to a 25 mL Hamilton syringe, and lowered into the lateral ventricle (DV 2.4 mm). After 7 days of

recovery, a-FMH (1 mg/mL in sterile saline) was administered at 1 mL per min for 5 min using an infusion pump. The needle was

left in place for one additional min after each infusion (Provensi et al., 2014).

Lipid Extraction and FractionationWe used a Bligh and Dyer procedure, modified as described (Astarita and Piomelli, 2009). Briefly, frozen tissues (z30-40 mg) were

weighed, transferred to glass vials and homogenized in cold methanol (2 mL) containing [2H4]-OEA, [2H4]-PEA, N-heptanoyl-PE and

heptadecanoic acid as internal standards. Lipids were extracted with chloroform (2mL) and washed with LC/MS-grade water (1 mL).

After centrifugation for 15 min at 2,850xg and 4�C, the organic phases were collected and transferred to a new set of glass vials. To

increase extraction efficiency, the aqueous fractions were extracted again with chloroform (1 mL) and the centrifugation step was

repeated. Both organic phases were pooled and dried under N2. Lipids were reconstituted in chloroform (2 mL) and the organic

extracts were fractionated by Silica Gel G column chromatography (60-A 230-400 mesh; Sigma-Aldrich, Milan, Italy). OEA, PEA

and fatty acids were eluted from the silica column with 2 mL of chloroform/methanol (9:1, v/v), N-acyl-PE species were eluted

with 2 mL of chloroform/methanol (1:1, v/v). The solvent was evaporated under N2 and lipids were reconstituted in methanol/

chloroform (0.1 mL; 9:1, v/v) and transferred to glass vials for LC/MS analyses.

LC/MS AnalysesWe measured OEA using a Xevo TQ UPLC-MS/MS system equipped with a reversed-phase BEH C18 column (2.1350 mm, 1.7 mm

particle size) (Waters, Milford, USA). The mobile phase consisted of 0.1% formic acid in water as solvent A and 0.1% formic acid in

acetonitrile as solvent B. A linear gradient was used: 0.0–0.5 min 20% B; 0.5–2.5 min 20 to 100% B; and 2.5–3.0 min maintained at

100% B. The column was reconditioned to 20% B for 1 min. Analysis time was 4 min and the injection volume was 5 mL. The mass

spectrometer was operated in the positive electrospray ionization (ESI) mode and quantifications were performed using the multiple

reaction monitoring (MRM) transition 326>62 for OEA, 330>66 for [2H4]-OEA, 300>62 for PEA and 304>66 for [2H4]-PEA. Capillary

voltage was 3 kV. Cone voltage and collision energy values were 25 V and 20 eV, respectively. Source temperature was 120�C.Desolvation gas and cone gas (nitrogen) flows were 800 L/h and 20 L/h, respectively. Desolvation temperature was 450�C. The limit

of quantification was 0.5 fmol on column.

N-acyl-PE species were identified and quantified using a Xevo TQ UPLC-MS/MS system equipped with a reversed-phase HSS

T3 C18 column (2.1350 mm, 1.8 mm particle size) (Waters). The mobile phase consisted of 10 mM ammonium formate (pH=5.0)

in acetonitrile/water (60:40, v/v) as solvent A and 10 mM ammonium formate (pH=5.0) in acetonitrile/isopropanol (10:90, v/v) as

solvent B. A linear gradient was used: 0.0–1.0 min 05% B; 1.0–7.0 min 0.5 to 100% B; and 7.0–7.9 min maintained at 100% B.

The column was reconditioned to 50% B for 1 min. The total analysis time was 9 min and the injection volume was 5 mL. The

mass spectrometer was operated in the positive ESI mode and quantification was performed using the MRM transitions

reported in Table S1. Capillary voltage was 3 kV. Cone voltage and collision energy values were 30 V and 25 eV, respectively. Source

Cell Metabolism 29, 91–102.e1–e5, January 8, 2019 e3

temperature was 120�C. Desolvation gas and cone gas (nitrogen) flows were 800 L/h and 20 L/h, respectively. Desolvation temper-

ature was 450�C. The limit of quantification was 0.5 fmoles on column.

We measured oleic acid using an Acquity UPLC system coupled with a Xevo TQ-MS triple quadrupole mass spectrometer.

Separation was carried out on aHSSC18 column (2.1 x 100mm, 1.7 mmparticle size) (Waters) at a flow rate of 0.4mL/min. Themobile

phase consisted of 20 mM ammonium acetate in water as solvent A and acetonitrile/isopropyl alcohol (1:1, v/v) as solvent B. A linear

gradient was used: 0.0–0.5 min 50% B; 0.5–3.5 min 50 to 100% B; 3.5–4.5 min maintained at 90% B; 4.5 to 5.5 min 90 to 100% B;

and maintained at 100% B for further 1.5 min. The column was reconditioned to 50% B for 1.5 min. Analysis time was 9 min and

the injection volume was 5 mL. The mass spectrometer was operated in the negative ESI mode and quantification was performed

using the MRM transition 281.2>281.2 and 269.2>269.2 for heptadecanoic acid (internal standard). Capillary voltage was 2.5 kV.

Cone voltage and collision energy were 25 V and 5 eV, respectively. Source temperature was 120�C. Desolvation gas and cone

gas (nitrogen) flows were 800 L/h and 50 L/h, respectively. Desolvation temperature was 400�C. The limit of quantification was

50 fmoles on column.

For histamine analysis, we transferred plasma samples (0.1 mL) to 1.5 mL plastic vials and precipitated proteins by addition of cold

acetonitrile (0.5 mL) containing [2H4]-histamine (100 nM) as internal standard. Samples were stirred for 1 min and centrifuged for

10 min at 3,000xg and 4�C. The supernatants were collected in glass vials, dried under N2, reconstituted in LC/MS-grade water

(0.1mL) and transferred to LC/MS vials. Separately, a calibration curve was prepared in histamine-depleted plasma (HDP) as follows.

HDP was prepared by adding activated charcoal (0.3 g) to 5 mL of mouse plasma and incubating the suspension at 37�C for 2 h with

stirring. The charcoal was removed by centrifugation for 10 min at 18,800xg. Authentic histamine was added to HDP at concentra-

tions ranging from 10 nM to 1000 nM along with [2H4]-histamine (100 nM). LC/MS analyses were conducted using a Xevo TQ

UPLC-MS/MS system equipped with an Acquity BEH HILIC column (2.13100 mm, 1.7 mm particle size) (Waters). The mobile phase

consisted of 0.5% formic acid in 25 mM ammonium formate as solvent A and acetonitrile as solvent B. A linear gradient was used:

0.0–0.5 min 98% B; 0.5–2.0 min 98% to 50% B; and 2.0–3.0 min maintained at 50% B. The column was reconditioned to 98% B for

1.5 min. Analysis time was 4.5 min and the injection volume was 5 mL. The mass spectrometer was operated in the positive ESI

mode and quantifications were performed using the MRM transitions 112 [M+H]+>95 [M-NH3+H]+ for histamine and 116>99 for

[2H4]-histamine. The capillary voltage was 2.8 kV. The cone voltage and collision energy values were 35 V and 15 eV, respectively.

Desolvation temperature was 450�C. Desolvation gas and cone gas (nitrogen) were 800 L/h and 50 L/h, respectively. The limit of

quantification was 50 fmoles on column.

Ketone Body Measurementsb-OHB was measured in plasma and liver samples using a colorimetric assay kit (Cayman Chemicals), following manufacturer’s

instructions.

Gene Expression AnalysesWe extracted total RNA from liver tissue using TRIzolTM reagent (Thermo Fisher Scientific, Monza, Italy) and quantified with the

PureLinkTM RNA Mini Kit (Thermo Fisher Scientific) as directed by the supplier. Samples were rendered genomic DNA-free by

treatment with DNase (PureLinkTM DNase, Thermo Fisher Scientific, Monza, Italy). cDNA was synthesized from 1 mg of total RNA

by using SuperScript VILOTM cDNA Synthesis Kit (Thermo Fisher Scientific) following manufacturer’s instructions. First-strand

cDNA was amplified using the iTaqTM Universal SYBR Green Supermix (Bio-Rad, Milan, Italy). The primer sequences were:

Acetyl-CoA acetyltransferase-1 (Acat-1) forward: 5’-GTCTGGCTAGTATTTGCAACG-3’; reverse: 5’-TTCAGCCGGTCACATGG-3’;

b-Hydroxy-b-methylglutaryl-CoA synthase-2 (Hmgcs-2) forward: 5’-TGGTTCAAGACAGGGCACACAGAAC-3’; reverse: 5’-AGAG

GAATACCAGGGCCCAACAAT-3’; Fibroblast growth factor 21 (Fgf21) forward: 5’-ATCAGGGAGGATGGAACAGTGG-3’; reverse:

5’-AGCTCCATCTGGCTGTTGGCAA-3’; Cytochrome P450, family 4, subfamily a, polypeptide 10 (Cyp4a10) forward: 5’-GCTACT

CAAGGCTTTCCAGCAG-3’; reverse: 5’-CCAGAACCATCTAGGAAAGGCAC-3’. Real-time PCR reactions were performed using

ViiA�7 Real-Time PCR detection system (Thermo Fisher Scientific). Thermal cycling conditions were: 50�C for 2 min followed by

an initial denaturation step at 95�C for 10 min, followed by 40 cycles, where each cycle was performed at 95�C for 15 s and at

60�C for 60 s. The Bestkeeper software (Pfaffl et al., 2004) was used to determine the expression stability and the geometric

mean of three different housekeeping genes [b-Actin, Hypoxanthine-guanine phosphoribosyltransferase (Hgprt) andGlyceraldehyde

phosphate dehydrogenase (Gapdh)]. The relative quantity of genes of interest was calculated by the 2-DDCt method (Livak and

Schmittgen, 2001). Data are reported as fold change relative to control groups.

ChemicalsWe purchased OEA, [2H4]-OEA, PEA and [2H4]-PEA from Cayman Chemicals (Ann Arbor, Michigan, USA). Fexofenadine, famotidine,

ciproxifan, heptadecanoic acid, fatty acids, histamine, and [2H4]-histamine (histamine-a,a, b, b-2H4) were from Sigma-Aldrich (Milan,

Italy). 1-stearoyl, 2-docosahexaenoyl-phosphatidylethanolamine N-heptanoyl (N-heptanoyl-PE) was synthesized in the lab as

described (Basit et al., 2016). a-FMH and JNJ7777120 were a generous gift from Dr. Nicholas Carruthers (Johnson & Johnson;

San Diego, USA).

e4 Cell Metabolism 29, 91–102.e1–e5, January 8, 2019

QUANTIFICATION AND STATISTICAL ANALYSIS

Datawere evaluated using Student t-test, One-way or Two-way ANOVA followed by Tukey’s or Bonferroni’s post hoc tests, as appro-

priate. Differences between groups were considered statistically significant if P < 0.05. Data were presented as mean ± SEM.

Significant outliers were excluded using the Grubbs’ test, considered statistically significant at P < 0.05. Statistical analyses were

performed using GraphPad Prism version 5.03 (GraphPad Software Inc., San Diego, CA, USA).

Cell Metabolism 29, 91–102.e1–e5, January 8, 2019 e5