Abstract
Reactive oxygen species exacerbate nonalcoholic steatohepatitis (NASH)
by oxidizing macromolecules; yet how they promote NASH remains poorly
understood. Here, we show that peroxidase activity of global hepatic
peroxiredoxin (PRDX) is significantly decreased in NASH, and palmitic
acid (PA) binds to PRDX1 and inhibits its peroxidase activity. Using
three genetic models, we demonstrate that hepatic PRDX1 protects
against NASH in male mice. Mechanistically, PRDX1 suppresses STAT
signaling and protects mitochondrial function by scavenging hydrogen
peroxide, and mitigating the oxidation of protein tyrosine phosphatases
and lipid peroxidation. We further identify rosmarinic acid (RA) as a
potent agonist of PRDX1. As revealed by the complex crystal structure,
RA binds to PRDX1 and stabilizes its peroxidatic cysteine. RA
alleviates NASH through specifically activating PRDX1’s peroxidase
activity. Thus, beyond revealing the molecular mechanism underlying PA
promoting oxidative stress and NASH, our study suggests that boosting
PRDX1’s peroxidase activity is a promising intervention for treating
NASH.
Subject terms: Metabolic disorders, Metabolic syndrome
__________________________________________________________________
Oxidative stress is closely linked with nonalcoholic steatohepatitis
(NASH). Here, the authors show that palmitic acid stimulates NASH by
inhibiting PRDX1 to increase oxidative stress, while rosmarinic acid
improves NASH by activating PRDX1 to reduce oxidative stress.
Introduction
Nonalcoholic fatty liver disease (NAFLD), defined as ectopic lipid
accumulation in hepatocytes without excessive alcohol consumption and
other causes like viral infections or drug abuse^[68]1, is the most
prevalent liver disease worldwide^[69]2 and essentially composed of two
forms: non-progressive form with simple steatosis and progressive form
(nonalcoholic steatohepatitis, NASH) that may advance to fibrosis,
cirrhosis and hepatocellular carcinoma (HCC)^[70]3. NASH is mostly
characterized by steatosis, oxidative stress, inflammation, and
hepatocyte ballooning with varying degrees of fibrosis^[71]3, and has
emerged as a risk factor of cardiovascular disease and type 2 diabetes
(T2D)^[72]4,[73]5. The pathogenesis of NASH is complex and still under
debate with different hypotheses^[74]6–[75]8. Among various pathogenic
factors, oxidative stress plays an essential part in
NASH^[76]3,[77]8,[78]9. For instance, oxidative stress is involved in
NASH pathogenesis through boosting inflammation^[79]7. Hydrogen
peroxide (H[2]O[2]) activates nuclear factor kappa B (NF-κB) and
stimulates the inflammatory response^[80]10. Increased reactive oxygen
species (ROS) and the resultant lipid peroxidation promote the
inflammatory response^[81]6. Although hepatocyte ROS have been shown to
profoundly promote NASH and liver fibrosis^[82]3,[83]11,[84]12, how
they are controlled remains to be elucidated.
Oxidative stress was initially defined as an imbalance between
pro-oxidant and anti-oxidant favoring the former; however, in the past
decades, this definition has been redefined and now includes the
transfer of oxidizing equivalents from a peroxidase to another target
protein that is known as ‘redox relay’^[85]13,[86]14. Thus, oxidative
stress exerts a tremendous impact both physiologically and
pathologically^[87]13. The anti-oxidant system consists of
non-enzymatic and enzymatic anti-oxidants including superoxide
dismutase (SOD), catalase, glutathione peroxidase, and peroxiredoxin
(PRDX)^[88]13,[89]15. A protective role for anti-oxidants in NASH has
been implicated in one study showing reduced glutathione and activities
of catalase and SOD in NASH patients^[90]16. The mammalian PRDX family
is composed of six peroxidases and guards against oxidative stress by
scavenging the majority of intracellular peroxides^[91]17–[92]20. Of
interest, numerous studies have demonstrated that PRDX1 is involved in
a variety of intracellular events and human diseases by functioning as
either a peroxidase or a chaperone^[93]17,[94]18,[95]21–[96]27. On one
hand, PRDX1 has been shown to protect against alcohol-induced liver
injury through its peroxidase activity^[97]27, or alleviate
inflammation and NASH through its chaperone activity^[98]25,[99]26. On
the other hand, PRDX1 also has been suggested to promote inflammation
and aggravate liver injury through its chaperone
activity^[100]23,[101]24. Thus far, the functional role of PRDX1 in
NASH through its peroxidase activity remains poorly understood.
In this study, we show that the peroxidase activity of global hepatic
PRDX is significantly decreased in NASH mouse models. Palmitic acid
directly binds to PRDX1 and reduces its peroxidase activity. We
demonstrate that hepatic PRDX1 protects against NASH through three
mouse models. By scavenging H[2]O[2] and mitigating the oxidation of
protein tyrosine phosphatases and lipid peroxidation, PRDX1 suppresses
STAT1 and STAT3 signaling and protects liver mitochondrial function.
Finally, we identify rosmarinic acid (RA) as a potent agonist of PRDX1
with a dissociation constant at nanomolar levels. As revealed by the
complex crystal structure, RA binds to PRDX1 and stabilizes its
peroxidation cysteine. RA treatment in WT mice alleviates NASH and
fibrosis, while it loses these beneficial effects in PRDX1 knockout
mice. Thus, boosting PRDX1’s peroxidase activity is a promising
therapeutic intervention for NASH.
Results
Decreased peroxidase activity of global hepatic PRDX in NASH
To examine whether the global PRDX peroxidase activity in the liver was
altered in NASH, we applied a classic Trx-TrxR-NADPH coupled
assay^[102]28,[103]29 where PRDX peroxidase activity is coupled to
NADPH oxidation via thioredoxin (Trx) and thioredoxin reductase (TrxR),
and accordingly can be evaluated indirectly by measuring NADPH
consumption rates (Supplementary Fig. [104]1a). Given that the majority
of intracellular peroxides are thought to be quenched by PRDX family
members^[105]18, the Trx-TrxR-NADPH coupled assay can be applied to
measure the global PRDX peroxidase activity. Using this assay, we
compared the global hepatic PRDX peroxidase activity in various NASH
mouse models induced by special diets with that in mice fed a normal
chow (NC) diet. High-fat diet (HFD) feeding for 18 weeks significantly
increased serum aspartate aminotransferase (AST) and alanine
aminotransferase (ALT) levels, which are indicative of liver
injury^[106]30 (Supplementary Fig. [107]1b), and hepatic H[2]O[2]
levels that were revealed by staining of HKPerox-Red, a selective
H[2]O[2] probe^[108]31 (Supplementary Fig. [109]1c).
As demonstrated by the reduced NADPH consumption rates, HFD
significantly decreased the global PRDX peroxidase activity in the
liver (Fig. [110]1a). HFD did not alter the protein levels of most PRDX
family enzymes, but significantly increased PRDX4 protein levels in the
liver (Fig. [111]1b). Interestingly, previous studies have demonstrated
that exposure to H[2]O[2] can hyperoxidize some PRDX members
(PRDX1-PRDX3) at their active cysteine to form sulfinic acid (SO2) or
sulfonic acid (SO3)^[112]17. Of note, formation of PRDX-SO2 causes a
reversible inactivation of PRDX peroxidase activity^[113]32, while
formation of PRDX-SO3 causes an irreversible inactivation of PRDX
peroxidase activity. Thus, we used an antibody (Anti-Peroxiredoxin-SO3,
ab16830, abcam) that recognizes both SO2 and SO3 forms of PRDX^[114]33
to detect PRDX hyperoxidation. Our observation of no difference in the
extent of hepatic PRDX hyperoxidation between NC and HFD groups
(Supplementary Fig. [115]1d) excluded that the observed decrease in the
global PRDX peroxidase activity after HFD results from hyperoxidation.
Fig. 1. Decreased peroxidase activity of global hepatic PRDX in NASH.
[116]Fig. 1
[117]Open in a new tab
a Peroxidase activity of global hepatic PRDX after normal chow (NC) or
high-fat diet (HFD) feeding. 8-week-old male C57BL/6 mice were fed a NC
or HFD for 18 weeks and liver samples were collected for biochemical
analyses. Global hepatic PRDX peroxidase activity was measured using a
classic Trx-TrxR-NADPH coupled assay. NADPH consumption was monitored
via absorbance at 340 nm (A[340]) in 15 min assay duration. Meanwhile,
the background activity was assessed without Trx and TrxR, but only
with H[2]O[2] and NADPH. To calculate the initial NADPH consumption
rate (initial rate) (A[340]/min/protein (g)) in the first 5 min, a
smooth curve was drawn through A[340] readings, and the initial rate
was calculated by performing a simple linear regression. Global PRDX
peroxidase activity was calculated by subtracting the background
activity (initial rate) from total activity (initial rate). n = 6 mice
per group. b Protein levels of hepatic PRDX family enzymes after HFD
(as in a) and quantitation. n = 6 mice per group; ns, no significance.
c Peroxidase activity of global hepatic PRDX after NC or western diet
(WD). 8-week-old male C57BL/6 mice were fed a NC or WD for 20 weeks and
global hepatic PRDX peroxidase activity was measured using a classic
Trx-TrxR-NADPH coupled assay. n = 6 mice per group. d Protein levels of
hepatic PRDX family enzymes after NC or WD feeding (as in c) and
quantitation. n = 6 mice per group; ns, no significance. e Peroxidase
activity of global hepatic PRDX after NC or methionine and choline
deficient diet (MCD). 8-week-old male C57BL/6 mice were fed a NC or MCD
for 5 weeks and global hepatic PRDX peroxidase activity was measured
using a classic Trx-TrxR-NADPH coupled assay. n = 6 mice per group. f
Protein levels of hepatic PRDX family enzymes after NC or MCD feeding
(as in e) and quantitation. n = 6 mice per group; ns, no significance.
All data are presented as means ± SEM. Unpaired and two-tailed
Student’s t test was performed for a–f.
We also assessed the global PRDX peroxidase activity in the liver in
two additional common experimental models of NASH, including NASH with
obesity induced by a western diet (WD) that is rich in fat, fructose,
and cholesterol, and NASH without obesity induced by a methionine and
choline deficient diet (MCD)^[118]7. WD feeding (20 weeks)
significantly decreased the global hepatic PRDX peroxidase activity and
protein levels of PRDX1 and PRDX4, but increased PRDX3 levels in the
liver (Fig. [119]1c, d). WD significantly increased hepatic H[2]O[2]
levels as shown by HKPerox-Red staining (Supplementary Fig. [120]1e),
and hepatic levels of hepatic malondialdehyde (MDA) (Supplementary
Fig. [121]1f), a marker for lipid peroxidation^[122]9, and serum levels
of ALT and AST (Supplementary Fig. [123]1g). WD feeding did not affect
the extent of hepatic PRDX hyperoxidation (Supplementary Fig.[124]1h).
We also observed that MCD feeding (5 weeks) significantly reduced the
global PRDX peroxidase activity in the liver (Fig. [125]1e) and protein
levels of PRDX1, PRDX5, and PRDX6, but significantly increased PRDX3
levels in the liver (Fig. [126]1f). MCD significantly increased hepatic
H[2]O[2] levels (Supplementary Fig. [127]1i), and hepatic levels of
hepatic MDA (Supplementary Fig. [128]1j) and serum levels of ALT and
AST (Supplementary Fig. [129]1k). MCD feeding had no effect on the
extent of hepatic PRDX hyperoxidation as indicated (Supplementary
Fig. [130]1l). Collectively, these results establish that the global
PRDX peroxidase activity in the liver is reduced in NASH, and suggest
that decreased PRDX peroxidase activity contributes to hepatic
oxidative stress and NASH pathogenesis.
PA binds to PRDX1 and inhibits its peroxidase activity
We next investigated potential causes of the decreased PRDX peroxidase
activity in NASH. Free fatty acids (FFA) and pro-inflammatory cytokines
such as interleukin 6 (IL-6) and interferons are elevated in
NASH^[131]3,[132]34. PA is the most common and abundant saturated FFA
with potent toxic effects on promoting NASH^[133]35. PA treatment
(1 hr) in HepG2 cells significantly decreased the global PRDX
peroxidase activity and increased the intracellular ROS levels
(Fig. [134]2a, b). In addition, treatment with PA at higher
concentration of (500 μM vs 250 μM) significantly increased H[2]O[2]
levels in HepG2 cells (Fig. [135]2c, d). Of note, PA treatment did not
alter PRDX protein levels (Supplementary Fig. [136]2a), or the extent
of PRDX hyperoxidation (Supplementary Fig. [137]2b), supporting that
the observed decrease in the global PRDX peroxidase activity is
directly induced by PA treatment.
Fig. 2. PA binds to PRDX1 and inhibits its peroxidase activity.
[138]Fig. 2
[139]Open in a new tab
a Global PRDX peroxidase activity in HepG2 cells after PA (250 μM)
treatment for 1 hr. Veh, vehicle. PA, palmitic acid. n = 5 biologically
independent samples. b ROS levels in HepG2 cells after PA treatment for
1 hr. Intracellular ROS were monitored by measuring the fluorescent
intensity of dichlorofluorescin (DCF) using a fluorometer. n = 4
independent experiments. c Representative images of HKPerox-Red
staining in HepG2 cells after treatment with Veh or PA at different
concentrations for 1 hr. Arrows denote the signals of HKPerox-Red
staining. n = 5 independent experiments. Scale bar, 50 μm. d
Quantification of fluorescence intensity of images from c. n = 5
independent experiments. e Binding affinity of sodium palmitate on
recombinant PRDX1 determined by SPR assay. Data were calculated from
three independent experiments. f Representative images showing the
increased thermal stabilization of PRDX1 after binding to PA and
quantification. n = 3 independent experiments. HepG2 cells were treated
with PA (250 μM) for 1 hr and then the cell lysate was collected for
the thermal shift assay. In brief, the cell lysate was divided into six
aliquots. One aliquot was used for input control and the other five
aliquots were heated at different temperatures as indicated for 3 min.
Finally, western blotting was carried out to detect PRDX1 stability. g
Inhibition of recombinant WT PRDX1’s peroxidase activity by sodium
palmitate at different concentrations as indicated. For more details,
please see methods section. Data were calculated from three independent
experiments. All the data are presented as means ± SEM. Unpaired and
two-tailed Student’s t test was performed for a, b, d, and f.
To test whether PA directly binds to PRDX1, we performed the surface
plasmon resonance (SPR) assay, where sodium palmitate was used due to a
poor solubility of PA. Sodium palmitate directly bound to recombinant
WT PRDX1, yielding a dissociation constant (K[D]) of 75.3 ± 4.5 μM
(Fig. [140]2e). Using the cellular thermal shift assay^[141]36, we also
observed the significantly increased thermal stabilization of PRDX1
upon PA binding in HepG2 cells (Fig. [142]2f). We further evaluated the
effect of sodium palmitate on recombinant WT PRDX1’s peroxidase
activity though the Trx-TrxR-NADPH assay. Sodium palmitate inhibited
PRDX1’s peroxidase activity in a dose dependent manner, with a maximal
inhibition rate at 46.6% (Fig. [143]2g).
To further evaluate the in vivo role of PA in NASH pathogenesis, we fed
mice a choline-deficient, amino acid-defined, HFD (CDAHFD) (another
common diet to induce NASH^[144]37) that contains different amounts of
PA (10% PA vs 20% PA) but same energy density (Supplementary
Table [145]3)^[146]38. High PA CDAHFD did not change body weight, but
significantly increased liver weight, serum levels of ALT, AST and TG,
and hepatic MDA contents (Supplementary Fig. [147]2c–g). In addition,
high PA CDAHFD significantly reduced the global hepatic PRDX peroxidase
activity, compared with low PA CDAHFD and NC groups (Supplementary
Fig. [148]2h). Histologically, we observed more severe phenotypes of
NASH and liver fibrosis in high PA CDAHFD-fed mice than low PA
CDAHFD-fed mice, as assessed by staining of hematoxylin and eosin
(H&E), Oil Red O, and smooth muscle actin alpha (α-SMA) (Supplementary
Fig. [149]2i). Collectively, these results suggest that PA promotes
NASH by inhibiting PRDX1’s peroxidase activity and increasing oxidative
stress.
PRDX1 knockout exacerbates NASH and liver fibrosis
To investigate potential impacts of PRDX1 in NASH pathogenesis, we
generated a PRDX1 knockout (Prdx1^-/-) mouse strain (Fig. [150]3a), and
observed the significantly reduced peroxidase activity of global
hepatic PRDX in Prdx1^-/- mice (Fig. [151]3b). After 6 weeks of WD
feeding, Prdx1^-/- mice had significantly higher body weights than
their wild-type (WT) littermates (Fig. [152]3c). Note that there was no
difference of food intake between WT and Prdx1^-/- mice (Fig. [153]3d),
suggesting that increased body weight of Prdx1^-/- mice could be owing
to decreased energy expenditure. Compared with WT mice, Prdx1^-/- mice
exhibited significantly decreased insulin sensitivity, as assessed with
intraperitoneal glucose tolerance test (IPGTT) and intraperitoneal
insulin tolerance test (IPITT) (Fig. [154]3e, f), as well as
significantly increased serum ALT and AST levels (Fig. [155]3g).
Prdx1^-/- mice showed significantly increased hepatic H[2]O[2] levels,
fat accumulation, and liver fibrosis as assessed by staining of
HKPerox-Red, H&E, Oil Red O, Sirius Red, and α-SMA (Fig. [156]3h–j).
Fig. 3. PRDX1 knockout increases NASH and liver fibrosis.
[157]Fig. 3
[158]Open in a new tab
a Representative images validating the efficiency of PRDX1 knockout in
Prdx1^-/- mice. This experiment was repeated for three times
independently. b Peroxidase activity of global hepatic PRDX in WT and
Prdx1^-/- mice. WT mice (n = 9); Prdx1^-/- mice (n = 8). c Body weight
of WT and Prdx1^-/- mice on WD. 8-week-old male mice were fed a WD for
20 weeks and their weekly body weights were monitored. n = 10 mice per
group. d Daily food intake of mice on WD (as in c). n = 10 mice per
group; ns, no significance. e Intraperitoneal glucose tolerance test
(IPGTT) in mice on WD (as in c) and area under the curve (AUC). n = 10
mice per group. f Intraperitoneal insulin tolerance test (IPITT) in
mice on WD (as in c) and AUC. n = 10 mice per group. g Circulating ALT
and AST levels (as in c). WT mice (n = 8); Prdx1^-/- mice (n = 10). h
Representative images of HKperox-Red staining in the liver and
quantitative analysis (as in c). Arrows denote the signals of
HKPerox-Red staining. Scale bar, 50 μm. n = 9 images from three mice
per group. i Representative images showing H&E and Oil Red O staining
in the liver after WD (as in c). n = 3 biologically independent mice.
Scale bars, 50 μm. j Representative images showing Sirius Red and α-SMA
staining in the liver (as in c). n = 3 biologically independent mice.
Scale bars, 50 μm. All data are presented as means ± SEM. Unpaired and
two-tailed Student’s t test was performed for b, d, AUC of e, AUC of f,
g, and h. Two-way ANOVA followed by Bonferroni’s test for multiple
comparisons was performed for c, e, and f.
To test whether hepatic PRDX1 confers protection against NASH, we
generated floxed Prdx1 (Prdx1^fl/fl) mice (Supplementary Fig. [159]3a),
and crossed them with Albumin-Cre mice^[160]39 (Alb-Cre;Prdx1^fl/fl) to
delete Prdx1 specifically from the liver (Supplementary Fig. [161]3b).
The peroxidase activity of global hepatic PRDX was significantly
reduced in Alb-Cre;Prdx1^fl/fl mice (Supplementary Fig. [162]3c).
When fed a WD (20 weeks), Alb-Cre;Prdx1^fl/fl mice showed no
differences from Prdx1^fl/fl control mice in body weight (Supplementary
Fig. [163]3d), food intake (Supplementary Fig. [164]3e), energy
expenditure (Supplementary Fig. [165]3f), and insulin sensitivity
(Supplementary Fig. [166]3g, h), but they uniquely increased steatosis
and liver fibrosis (Supplementary Fig. [167]3i, j). In agreement with
these phenotypes, the expression of several hepatic genes related to
inflammation and fibrosis (e.g., Cd11b, Col1a1, Col3a1, Pdgfb, and
Pdgfra) was significantly increased in Alb-Cre;Prdx1^fl/fl mice
(Supplementary Fig. [168]3k).
When fed a MCD (5 weeks), Alb-Cre;Prdx1^fl/fl mice showed similar
hepatic fat accumulation to Prdx1^fl/fl control mice (Supplementary
Fig. [169]3i), but exhibited severe liver fibrosis (Supplementary
Fig. [170]3m), and significantly increased the expression levels of
inflammatory or fibrotic genes (e.g, F4/80, Col3a1, Pdgfb, and Pdgfra)
in the liver (Supplementary Fig. [171]3n). We also observed significant
increases in H[2]O[2] levels and the extent of lipid peroxidation in
the liver of Alb-Cre;Prdx1^fl/fl mice (Supplementary Fig. [172]3o, p).
Collectively, these results demonstrate that PRDX1 knockout exacerbates
NASH and liver fibrosis.
PRDX1 overexpression ameliorates NASH and liver fibrosis
To further understand how PRDX1 influences NASH, we made a PRDX1
overexpression (Prdx1^OE/OE) mouse line (Supplementary Fig. [173]4a,
b). Prdx1^OE/OE mice showed significantly increased peroxidase activity
of global hepatic PRDX (Fig. [174]4a).
Fig. 4. PRDX1 overexpression ameliorates NASH and liver fibrosis.
[175]Fig. 4
[176]Open in a new tab
a Peroxidase activity of global hepatic PRDX in WT and Prdx1^OE/OE
mice. WT mice (n = 5); Prdx1^OE/OE mice (n = 7). b Body weight of WT
and Prdx1^OE/OE mice on WD. 8-week-old male mice were fed a WD and
their body weights were monitored weekly. WT mice (n = 6); Prdx1^OE/OE
mice (n = 5). c Daily food intake of mice on WD (as in b). WT mice
(n = 6); Prdx1^OE/OE mice (n = 5). ns, no significance. d Energy
expenditure (kcal) of mice on WD (as in b). WT mice (n = 6);
Prdx1^OE/OE mice (n = 5). e Locomotion activity of mice on WD (as in
b). WT mice (n = 6); Prdx1^OE/OE mice (n = 5). ns, no significance. f
Fasting blood glucose levels of mice on WD (as in b). WT mice (n = 6);
Prdx1^OE/OE mice (n = 5). g IPITT in mice on WD (as in b). WT mice
(n = 6); Prdx1^OE/OE mice (n = 5). h Serum ALT and AST levels in mice
on WD (as in b). WT mice (n = 6); Prdx1^OE/OE mice (n = 5). i Hepatic
H[2]O[2] levels in WD-fed mice (as in b). n = 5 mice per group. j
Hepatic MDA levels in WD-fed WT and Prdx1^OE/OE mice (as in b). WT mice
(n = 6); Prdx1^OE/OE mice (n = 5). k Representative images showing H&E
and Oil Red O staining of liver after WD (as in b). n = 3 biologically
independent mice. Scale bars, 50 μm. l Representative images from three
mice per group showing Sirius Red and α-SMA staining of liver after WD
(as in b). n = 3 biologically independent mice. Scale bars, 50 μm. m
mRNA expression of hepatic genes after WD (as in b). WT mice (n = 6);
Prdx1^OE/OE mice (n = 5). All the data are presented as means ± SEM.
Unpaired and two-tailed Student’s t test was performed for a, c, d, e,
f, h, i, j, and m. Two-way ANOVA followed by Bonferroni’s test for
multiple comparisons was performed for b and g.
After 8 weeks of WD feeding, Prdx1^OE/OE mice had significantly less
body weights (Fig. [177]4b) but no difference in food intake than WT
mice (Fig. [178]4c), suggesting increased energy expenditure in
Prdx1^OE/OE mice. Indeed, Prdx1^OE/OE mice showed significantly
increased energy expenditure at night time (Fig. [179]4d), but similar
locomotion activity compared with WT mice (Fig. [180]4e). Prdx1^OE/OE
mice showed significantly improved insulin sensitivity, as demonstrated
by lower fasting glucose levels and significantly improved insulin
tolerance compared to WT mice (Fig. [181]4f, g), and showed significant
reductions in serum ALT and AST levels (Fig. [182]4h), hepatic H[2]O[2]
levels (Fig. [183]4i, and Supplementary Fig. [184]4c), and the extent
of hepatic lipid peroxidation (Fig. [185]4j). In addition, lipid
accumulation and fibrosis in the liver were markedly reduced in
Prdx1^OE/OE mice compared to WT mice (Fig. [186]4k, l). In line with
these phenotypes, numerous pro-inflammatory or fibrotic genes (e.g.,
Mcp-1, F4/80, Cd11b, Tnf-α, Il-6, Col1a1, Col3a1, Pdgfa and Pdgfra)
were significantly downregulated in the livers of Prdx1^OE/OE mice
(Fig. [187]4m).
When fed a MCD, Prdx1^OE/OE mice showed similar body weight
(Supplementary Fig. [188]4d), and fat accumulation in the liver
(Supplementary Fig. [189]4e), but significantly reduced liver fibrosis
(Supplementary Fig. [190]4f), the levels of hepatic H[2]O[2] and MDA
(Supplementary Fig. [191]4g, h), and the expression of hepatic
pro-inflammatory and fibrotic genes (Mcp-1, Il-1b, Col1a1, and Col3a1)
(Supplementary Fig. [192]4i). Together, these results demonstrate that
PRDX1 overexpression protects against NASH and liver fibrosis.
PRDX1 suppresses hepatic STAT1 and STAT3 signaling
Protein tyrosine phosphatases (PTPs) inhibit JAK-STAT
signaling^[193]40. Previous studies have shown that obesity-associated
ROS oxidize and inactivate PTPs (e.g., PTP1B and T cell protein
tyrosine phosphatase (TCPTP)), consequently increasing hepatic STAT
signaling and promoting NAFLD^[194]41,[195]42. Consistently, we
observed that hepatic PTPs’ oxidation (Fig. [196]5a, b), and hepatic
STAT1 and STAT3 phosphorylation (Fig. [197]5c, d) were drastically
increased in both WD- and MCD-induced NASH mouse models. Further,
H[2]O[2] treatment (30 min) in HepG2 cells dose-dependently increased
PTPs’ oxidization (Supplementary Fig. [198]5a), and STAT1 and STAT3
phosphorylation (Supplementary Fig. [199]5b). Together, these results
suggest that by oxidizing and inactivating PTPs, H[2]O[2] increases
STAT signaling and promotes NASH.
Fig. 5. PRDX1 suppresses hepatic STAT1 and STAT3 phosphorylation.
[200]Fig. 5
[201]Open in a new tab
a Western blotting of the oxidation of hepatic protein tyrosine
phosphatases (PTPs) in WT mice after NC or WD for 20 weeks. n = 4 mice
per group. b Western blotting of the oxidation of hepatic PTPs in WT
mice after NC or MCD for 2 weeks. n = 4 mice per group. c Western
blotting and quantitation of hepatic STAT1 and STAT3 phosphorylation
(as in a). n = 4 mice per group. d Western blotting and quantitation of
hepatic STAT1 and STAT3 phosphorylation (as in b). n = 4 mice per
group. e Western blotting and quantitation of hepatic STAT1 and STAT3
phosphorylation in MCD-fed WT and Prdx1^-/- mice. n = 4 mice per group.
f Western blotting and quantitation of hepatic STAT1 and STAT3
phosphorylation in WD-fed WT and Alb-Cre;Prdx1^fl/fl mice. n = 3 mice
per group. g Western blotting and quantitation of hepatic STAT1 and
STAT3 phosphorylation in WD-fed WT and Prdx1^OE/OE mice. n = 4 mice per
group. All the data are presented as means ± SEM. Unpaired and
two-tailed Student’s t test was performed for c–g.
IL-6 and IFN-γ are known to stimulate STAT signaling^[202]43. We
questioned if increased H[2]O[2] levels promote IL-6- and
IFN-γ-stimulated STAT signaling. Interestingly, treatment with either
IL-6- or IFN-γ in HepG2 cells significantly increased intracellular
H[2]O[2] levels (Supplementary Fig. [203]5c, d), and phosphorylation of
STAT1 and STAT3 (Supplementary Fig. [204]5e, f), which were drastically
reduced by pretreatment with a potent anti-oxidant (N-acetylcysteine
(NAC))^[205]44.
To study how PRDX1 regulates STAT signaling, we generated PRDX1
knockout HepG2 cells (Supplementary Fig. [206]5g), which showed
significantly reduced global PRDX peroxidase activity (Supplementary
Fig. [207]5h), and significantly increased intracellular H[2]O[2]
levels as well as extent of lipid peroxidation (Supplementary
Fig. [208]5i–k). In addition, the oxidation of PTPs (Supplementary
Fig. [209]5l), and IL-6- or IFN-γ-induced phosphorylation of STAT1 and
STAT3 (Supplementary Fig. [210]5m, n), all were drastically increased
in PRDX1 knockout HepG2 cells compared to WT HepG2 cells. Consistently,
PRDX1 knockout significantly increased hepatic STAT1 and STAT3
phosphorylation in MCD- and WD-induced NASH (Fig. [211]5e, f, and
Supplementary Fig. [212]5o). In contrast, PRDX1 overexpression
significantly reduced the phosphorylation of hepatic STAT1 and STAT3 in
WD-induced NASH (Fig. [213]5g). These results collectively indicate
that PRDX1 suppresses STAT signaling by scavenging H[2]O[2] and
mitigating the oxidation of PTPs.
To further support that PRDX1 suppresses STAT signaling, we performed
RNA sequencing of liver samples from WT and Alb-Cre;Prdx1^fl/fl mutant
mice fed a MCD. Gene-set enrichment analysis (GSEA) and KEGG pathway
enrichment analysis revealed enrichments for genes related to JAK-STAT
signaling pathway in Alb-Cre;Prdx1^fl/fl mutant mice (Supplementary
Fig. [214]6a, b). Intriguingly, the expression of numerous genes in
JAK-STAT signaling including some fibrotic genes such as Pdgfb and
Pdgfra was significantly increased in Alb-Cre;Prdx1^fl/fl mutant mice
(Supplementary Fig. [215]6c).
PRDX1 protects liver mitochondrial function
Liver mitochondrial dysfunction, which could be caused by hepatic lipid
peroxidation^[216]45, is a driving force of NASH^[217]46. Using an
Oroboros Oxygraph-2k (O2k) system, one clinical study showed that liver
mitochondrial function was compromised in NASH patients with increased
hepatic H[2]O[2] levels and lipid peroxidation, which mainly was
revealed by a significant increase in the leaking control ratio (LCR)
and a significant decrease in the respiratory control ratio
(RCR)^[218]47.
Using a similar O2k approach, we investigated the function of liver
mitochondria isolated from different NASH mouse models. Both WD (20
weeks) and MCD (5 weeks) significantly reduced the RCR (Fig. [219]6a
and Supplementary Fig. [220]7a) and increased the LCR in the liver
mitochondria of WT mice (Fig. [221]6b and Supplementary Fig. [222]7b).
In addition, citrate synthase activity (CSA) was significantly
increased in both NASH models (Fig. [223]6c and Supplementary
Fig. [224]7c), which is indicative of mitochondrial
dysfunction^[225]48. Of interest, mitochondrial oxygen (O[2]) flux (per
CSA) was significantly increased in the liver of MCD-fed mice
(Supplementary Fig. [226]7d), while no change of liver mitochondrial
O[2] flux was observed in WD-fed mice compared to NC-fed mice
(Fig. [227]6d). We also detected a significant increase in the extent
of lipid peroxidation in the liver mitochondria of WD- and MCD-fed mice
(Fig. [228]6e and Supplementary Fig. [229]7e). These results
collectively suggest that lipid peroxidation-induced mitochondrial
dysfunction contributes to the development and progression of NASH
(with or without obesity).
Fig. 6. PRDX1 protects liver mitochondrial function.
[230]Fig. 6
[231]Open in a new tab
a Hepatic respiratory control ratio (RCR) of mice on NC or WD.
8-week-old male C57BL/6 mice were fed a NC or WD for 20 weeks before
their liver mitochondria were isolated for O2k analyses. n = 3 mice per
group. b Hepatic leaking control ratio (LCR) (as in a). n = 3 mice per
group. c Hepatic citrate synthase activity (CSA) (as in a). n = 3 mice
per group. d O[2] flux in isolated liver mitochondria (as in a). n = 3
mice per group; ns, no significance. Mal, malate; Glu, glutamate; Suc,
succinate; Cyt, cytochrome c; Ccc, cccp. ns, no significance by
unpaired Student’s t test. e MDA concentration in the liver
mitochondria isolated from mice after NC or WD (as in a). n = 3 mice
per group. f Hepatic RCR of WT and Prdx1^OE/OE mice. 8-week-old male WT
and Prdx1^OE/OE mice were fed a WD for 20 weeks before their liver
mitochondria were isolated for O2k analyses. WT mice (n = 6);
Prdx1^OE/OE mice (n = 7). g Hepatic LCR (as in f). WT mice (n = 6);
Prdx1^OE/OE mice (n = 7). h Hepatic CSA (as in f). WT mice (n = 6);
Prdx1^OE/OE mice (n = 7). i O[2] flux in isolated liver mitochondria
(as in f). WT mice (n = 6); Prdx1^OE/OE mice (n = 7). j MDA
concentration in the liver mitochondria isolated from WT and
Prdx1^OE/OE mice after WD. n = 5 mice per group. All the data are
presented as means ± SEM. Unpaired and two-tailed Student’s t test was
performed for a–j.
To test whether PRDX1 protects liver mitochondrial function by
mitigating lipid peroxidation, we performed O2k analyses in Prdx1^OE/OE
and WT mice fed a WD (20 weeks) or a MCD (5 weeks) to induce NASH.
Compared to WT mice, Prdx1^OE/OE mice showed improved mitochondrial
function reflected by significantly increased RCR (Fig. [232]6f) and
significantly decreased LCR (Fig. [233]6g) after WD feeding. Hepatic
CSA tended to be significantly reduced in Prdx1^OE/OE mice (p = 0.053)
(Fig. [234]6h), while no difference in liver mitochondrial O[2] flux
was observed between WT and Prdx1^OE/OE mice (Fig. [235]6i). In
addition, the extent of lipid peroxidation was significantly reduced in
liver mitochondria of Prdx1^OE/OE mice (Fig. [236]6j).
We also observed a significant increase of RCR (Supplementary
Fig. [237]7f) and a significant decrease of LCR (Supplementary
Fig. [238]7g) in Prdx1^OE/OE mice compared with WT mice after MCD
feeding, although there was no difference in CSA and liver
mitochondrial O[2] flux between these two genotypes (Supplementary
Fig. [239]7h, i). Further, lipid peroxidation levels were significantly
reduced in the liver mitochondria of Prdx1^OE/OE mice (Supplementary
Fig. [240]7j). Together, these results support that beyond suppressing
STAT signaling, PRDX1 protects liver mitochondrial function by
scavenging H[2]O[2] and mitigating lipid peroxidation in the liver
mitochondria.
Identification of rosmarinic acid as an agonist of PRDX1
Our results collectively suggest that boosting PRDX1’s peroxidase
activity is a potential way in combating NASH. Through protein thermal
shift assay (PTS) based compound library screening and peroxidase
activity assay based validation, we identified rosmarinic acid (RA) as
a highly active agonist of PRDX1 (Fig. [241]7a and Supplementary
Fig. [242]8a). The half maximal concentration of RA for activating
PRDX1’s peroxidase activity is 253.1 ± 49.0 nM (Fig. [243]7b), while
the K[D] of RA with PRDX1 is 375.7 ± 2.5 nM as revealed by SPR
(Fig. [244]7c).
Fig. 7. Identification of rosmarinic acid as an agonist of PRDX1.
[245]Fig. 7
[246]Open in a new tab
a Identification of rosmarinic acid (RA) as a potential agonist of
PRDX1. Protein thermal shift assay (PTS) was used to identify 6 hits
and RA shows the highest efficacy in activating PRDX1’s peroxidase
activity as reflected by in vitro peroxidase activity assay. For more
details, please see the Methods section. Hits are marked as red dots,
while others are shown as black dots. Blue dashed lines represent the
same positive values of ΔT[m]D and ΔT[m]B. ΔT[m]D, derivative delta
melting temperature; ΔT[m]B, Boltzmann delta melting temperature; LA,
lawsone; MO, morine; SB, Salvianolic acid B; EG, Epigallocatechin
Gallate; PH, Phloracetophenone. b Half maximal concentration of RA for
activating WT PRDX1’s peroxidase activity. For a and b, data are
presented as means ± SEM from three independent experiments. c Binding
affinity of RA with WT PRDX1 by SPR assay. The dissociation constant
(K[D]) is shown as means ± SEM from duplicate experiments. d Unbiased
F[O]-F[C] density map contoured at 2.5σ of RA in RA-PRDX1^C52SC83S
(aa1-175) complex structure. RA is shown in yellow stick. Maps are
shown in blue nets. e 2F[O]-F[C] density map of RA and neighboring
residues of PRDX1 in RA- PRDX1^C52SC83S (aa 1-175) complex structure.
RA is shown in yellow stick. Maps are shown in gray nets. Waters are
shown in red spheres. Chain A and chain B are shown in green and navy
cartoon, respectively. Residues near RA are also shown as sticks. f
Complex crystal structure showing that RA binds at the peroxidatic site
of PRDX1^C52SC83S (aa 1-175). The peroxidatic site is highlighted in
salmon stick. RA, Chain A and Chain B are shown as in (e). g
Electrostatic potential of RA-PRDX1 ^C52SC83S (aa 1-175) complex
crystal structure and RA binding site. The interior of RA’s binding
site is electronegative (colored in blue), while the exterior is
electropositive (colored in red). h Hydrogen bond network formed
between RA and residues in the binding pocket. Residues around 5 Å of
RA are shown in stick. Waters are shown in red spheres. Hydrogen bonds
are shown in magenta dashed lines.
As a typical 2-Cys peroxidase, PRDX1 contains a peroxidatic cysteine
(C52) and a resolving cysteine (C173)^[247]17. To better understand the
mechanism of PRDX1 activation by RA, we solved a complex structure of
RA with PRDX1 variant, PRDX1^C52SC83S (aa1-175) (Supplementary
Table [248]1), where both C52 and C83 residues were mutated to serine,
and the C-terminus (aa 176-199) of WT PRDX1 protein was truncated,
given that only PRDX1^C52SC83S (aa1-175) variant could achieve
repeatable and high-resolution crystals after crystallization screening
with WT and different PRDX1 variants. F[O]-F[C] (Fig. [249]7d) and
2F[O]-F[C] (Fig. [250]7e) maps of RA and PRDX1^C52SC83S (aa 1-175) are
intact, indicating that the binding mode of RA in the complex structure
is reliable.
The complex structure demonstrates that molecular scaffold of RA
interacts with Chain A and Chain B of PRDX1, and RA binds at the
peroxidatic active site of PRDX1 (Fig. [251]7f). Electrostatic
potential map shows that the interior of RA binding site (peroxidatic
site) is electronegative, whereas the exterior is electropositive
(Fig. [252]7g). Overall, twelve hydrogen bonds between RA and
PRDX1^C52SC83S (aa 1-175) form a hydrogen bond network, thereby
stabilizing the residues in the binding site (Fig. [253]7h). Of
particular note, RA’s carboxyl group and carbonyl group form hydrogen
bonds with T49 and F50 of Region I, and V51 and S52 of the C[P] loop
(peroxidatic cysteine-containing loop) of PRDX1 (Supplementary
Fig. [254]8b). It has been suggested that salt-bridged hydrogen bonds
formed between the peroxidatic cysteine and conserved R128 stabilize
the active site and promote H[2]O[2] binding^[255]49. Thus, hydrogen
bonds between R128 and S52 bridged by RA’s carboxyl group help to
stabilize the peroxidatic cysteine (Supplementary Fig. [256]8c). Taken
together, we propose that the hydrogen bond network formed between RA
and the active site of PRDX1 helps to activate PRDX1’s peroxidase
activity.
We next validated the specificity of RA for PRDX1. RA treatment
(30 min) significantly increased the peroxidase activity of global PRDX
in WT HepG2 cells (Supplementary Fig. [257]8d); in contrast, RA
treatment in PRDX1 knockout HepG2 cells had no effect on PRDX
peroxidase activity (Supplementary Fig. [258]8e). Likewise, RA
treatment in WT mice significantly increased the peroxidase activity of
global hepatic PRDX (Supplementary Fig. [259]8f), but did not alter
hepatic PRDX peroxidase activity in PRDX1 knockout mice (Supplementary
Fig. [260]8g).
RA is a natural compound derived from plants with anti-oxidant and
anti-inflammatory activities^[261]50, and has shown hepatoprotective
effects^[262]51,[263]52; Consistently, RA treatment significantly
inhibited IL-6-induced ROS increase (Supplementary Fig. [264]8h), and
LPS-induced expression of pro-inflammatory cytokines (IL-6 and IL-1β)
in primary mouse hepatocytes (Supplementary Fig. [265]8i, j).
Interestingly, we found that RA dose-dependently reduced the levels of
recombinant WT PRDX1 hyperoxidation in vitro (Supplementary
Fig. [266]8k), as well as LPS-stimulated PRDX hyperoxidation in HepG2
cells (Supplementary Fig. [267]8l). Given that PRDX1 hyperoxidation
suppresses its peroxidase activity^[268]17, these data suggest that RA
could activate PRDX1’s peroxidase activity partially, if not
completely, by reducing its hyperoxidation. Collectively, these results
demonstrate that RA is a highly potent and specific agonist of PRDX1
with both anti-oxidant and anti-inflammatory activities.
RA treatment alleviates NASH and liver fibrosis
We next evaluated RA activity in WD-induced NASH. RA treatment
significantly increased the peroxidase activity of global PRDX in the
liver of WD-fed WT mice (Supplementary Fig. [269]9a). Although RA
treatment did not change body weight, energy expenditure and locomotion
activity (Supplementary Fig. [270]9b–d), it significantly improved
glucose intolerance (Supplementary Fig. [271]9e) and insulin
sensitivity (Supplementary Fig. [272]9f). In addition, RA treatment
effectively reduced the levels of hepatic H[2]O[2] and lipid
peroxidation (Fig. [273]8a, b, and Supplementary Fig. [274]9g), and
liver fibrosis as shown by the staining of Sirius Red and α-SMA
(Fig. [275]8c), though RA treatment did not alter fat accumulation in
the liver as shown by Oil Red O staining (Supplementary Fig. [276]9h).
Consistent with these phenotypes, we observed that in RA-treated mice
there was a significant reduction in the expression of numerous
pro-inflammatory or fibrotic genes (e.g., Mcp-1, Tnf-α, F4/80, Cd11b,
Cd11c, Col1a1, Col3a1, Pdgfa, Pdgfra and Pdgfb) (Fig. [277]8d), and in
hepatic STAT1 and STAT3 phosphorylation (Fig. [278]8e). Of note, RA
treatment significantly reduced the levels of PRDX hyperoxidation in
the liver (Supplementary Fig. [279]9i). In addition, RA treatment
improved the liver mitochondrial coupling and respiratory efficiency as
indicated by significantly reduced LCR, but did not alter CSA levels or
mitochondrial O[2] flux (Fig. [280]8f, g, and Supplementary
Fig. [281]9j, k).
Fig. 8. RA treatment alleviates NASH and liver fibrosis.
[282]Fig. 8
[283]Open in a new tab
a Hepatic H[2]O[2] levels in WT mice treated with WD and RA or vehicle.
8-week-old male WT mice were fed a WD and concurrently received a daily
intraperitoneal injection of either vehicle or RA (30 mg/kg) for 20
weeks. n = 6 mice per group. b Hepatic lipid peroxidation (as in a).
MDA levels were measured using a lipid peroxidation MDA assay kit.
n = 6 mice per group. c Representative images from three mice per group
showing Sirius Red and α-SMA staining of liver (as in a). n = 3
biologically independent mice. Scale bars, 50 μm. d mRNA expression of
hepatic genes (as in a). n = 6 mice per group. e Western blotting and
quantitation of hepatic STAT1 and STAT3 phosphorylation (as in a).
n = 4 mice per group. f Hepatic RCR. 8-week-old male WT mice were fed a
WD and concurrently received a daily intraperitoneal injection of
either vehicle or RA (30 mg/kg) for 20 weeks before liver mitochondria
were isolated for O2k analyses. n = 6 mice per group. ns, no
significance. g Hepatic LCR (as in f). n = 6 mice per group. h
Representative images showing Sirius Red and α-SMA staining in the
liver. 8-week-old male WT mice were fed a MCD and concurrently received
a daily intraperitoneal injection of vehicle or RA (30 mg/kg) for two
weeks. n = 3 biologically independent mice. Scale bars, 50 μm. i
Western blotting and quantitation of hepatic STAT1 and STAT3
phosphorylation (as in h). n = 4 mice per group. j mRNA expression of
hepatic genes (as in h). n = 5 mice per group. k Hepatic RCR.
8-week-old male WT mice were fed a MCD and concurrently received a
daily intraperitoneal injection of vehicle or RA for two weeks before
their liver mitochondria were isolated for O2k analyses. n = 5 mice per
group. l Hepatic LCR (as in k). n = 5 mice per group. All the data are
presented as means ± SEM. Unpaired and two-tailed Student’s t test was
performed for a–l, except c and h.
We also evaluated RA activity in MCD-induced NASH. RA treatment
significantly increased the peroxidase activity of global PRDX in the
liver of MCD-fed WT mice (Supplementary Fig. [284]9l). Although RA
treatment did not alter fat accumulation in the liver (Supplementary
Fig. [285]9m), it effectively reduced liver fibrosis (Fig. [286]8h),
hepatic STAT1 and STAT3 phosphorylation (Fig. [287]8i), and expression
of several inflammatory or fibrotic genes in the liver (e.g., Cd11c,
Col1a1, Col3a1, Pdgfb, and Pdgfra) (Fig. [288]8j). In addition, RA
treatment improved the liver mitochondrial function as revealed by
significantly decreased LCR and increased O[2] flux in the liver
mitochondria (Fig. [289]8k, l, and Supplementary Fig. [290]9n, o).
In contrast, RA treatment in WD-fed Prdx1^-/- mice did not change
hepatic H[2]O[2] levels and the extent of lipid peroxidation
(Supplementary Fig. [291]10a, b), nor mitigated NASH symptoms
(Supplementary Fig. [292]10c) and hepatic STAT1 and STAT3
phosphorylation (Supplementary Fig. [293]10d). In addition, RA
treatment did not improve liver mitochondria function as assessed with
O2k system (Supplementary Fig. [294]10e–h). Together, these results
indicate that RA protects against WD-induced NASH and fibrosis by
specifically activating PRDX1.
Collectively, through specifically activating PRDX1’s peroxidase
activity and reducing hepatic H[2]O[2] levels, RA treatment suppresses
hepatic STAT1 and STAT3 signaling activity, protects liver
mitochondrial function, and ultimately alleviates NASH and liver
fibrosis.
PRDX1 peroxidase dead mutant (PRDX1^Cys52Ser) confers resistance to NASH by
increasing the Hippo pathway
PRDX1 shows both anti-oxidative (peroxidase) and pro-inflammatory
(molecular chaperone) activities^[295]24,[296]27. Interestingly, one
previous study performed in cultured cells has implied that the
peroxidatic cysteine 52 (cys52) of PRDX1 could underlie its
pro-inflammatory activity^[297]53. To investigate whether PRDX1 cys52
mediates its pro-inflammatory activity in vivo, we recently generated a
PRDX1 mutant mouse model (PRDX1^Cys52Ser), where PRDX1 cys52 was
mutated to ser52 and consequently PRDX1’s peroxidase activity was
impaired^[298]54. Surprisingly, PRDX1^Cys52Ser mice showed robust
resistance to diet-induced NASH^[299]54. These findings are potentially
inconsistent with a protective role of PRDX1 against NASH through its
peroxidase activity as demonstrated in this study.
We sought to understand how PRDX1^Cys52Ser mutant confers resistance to
NASH although impairing PRDX1’s peroxidase activity. We fed WT and
PRDX1^Cys52Ser mice a CDAHFD and analyzed NASH phenotypes. Consistent
with our previously observed resistance of PRDX1^Cys52Ser mice to
either WD- or MCD-induced NASH^[300]54, PRDX1^Cys52Ser mice showed
significantly reduced hepatic inflammation and robust resistance to
NASH compared with WT mice after CDAHFD (Supplementary Fig. [301]11).
Next, using immunoprecipitation (IP) combined with mass spectrometry we
identified PPP1ca as a protein preferentially binding PRDX1^Cys52Ser
over WT PRDX1 (Supplementary Fig. [302]12a and b), which was further
confirmed by Co-IP (Supplementary Fig. [303]12c and d).
Numerous studies have demonstrated that as key downstream effectors of
the Hippo pathway, YAP and TAZ (YAP/TAZ) stimulate NASH and liver
fibrosis by promoting hepatic inflammation^[304]55–[305]60. In
addition, phosphorylation of YAP/TAZ leads to inhibition of their
activities^[306]55, and PPP1ca has been shown to dephosphorylate and
activate TAZ^[307]61. These together led us to postulate that
PRDX1^Cys52Ser mutant confers resistance to NASH by binding PPP1ca and
blocking its phosphatase activity, consequently increasing TAZ
phosphorylation and suppressing its activity. In line with this
hypothesis, the phosphorylation levels of both PPP1ca and TAZ were
significantly increased in the liver of CDAHFD-fed PRDX1^Cys52Ser mice
(Supplementary Fig. [308]13a). Note that PPP1ca phosphorylation
inhibits its phosphatase activity^[309]62,[310]63. These results
indicate that PRDX1^Cys52Ser mutant increases the Hippo pathway, which
was also evidenced by enrichment of genes related to the Hippo pathway
as revealed by RNA-Seq, and significantly decreased expression of a
number of YAP/TAZ downstream genes in the liver of CDAHFD-fed
PRDX1^Cys52Ser mice (Supplementary Fig. [311]13b, c). In addition, we
also observed significantly reduced phosphorylation of PPP1ca and TAZ
(Supplementary Fig. [312]14a), enrichment of genes related to the Hippo
pathway (Supplementary Fig. [313]14b), and significantly decreased
expression of several YAP/TAZ downstream genes in the liver of WD-fed
PRDX1^Cys52Ser mice compared with WD-fed WT mice (Supplementary
Fig. [314]14c). Taken together, these data suggest that PRDX1^Cys52Ser
mutation increases the Hippo pathway (TAZ inhibition) by increasing
PPP1ca binding and phosphorylation and consequently suppressing PPP1ca
phosphatase activity, a protective effect that outweighs loss of
PRDX1’s peroxidase activity and ultimately reduces hepatic inflammation
and ameliorates NASH.
Discussion
This study provides evidence that decreased peroxidase activity of
hepatic PRDX contributes to hepatic oxidative stress and promotes NASH.
Substantial evidence has demonstrated an intimate relationship between
ROS and NASH^[315]8. Nevertheless, research attention has been mainly
focused on ROS generation in the liver^[316]64–[317]66, especially in
non-hepatocytes^[318]67–[319]69. In contrast, how ROS clearance by
anti-oxidants influences NASH remains poorly understood^[320]70,
although several anti-oxidants have been shown to be reduced in NASH
patients^[321]16,[322]47. We demonstrate that decreased global hepatic
PRDX peroxidase activity accounts, at least partly, for increased
hepatic H[2]O[2] levels and NASH progression based on in vitro and in
vivo evidence. In line with our findings, previous studies have
suggested a protective role for other PRDX family enzymes including
PRDX4, PRDX5 and PRDX6 against NAFLD or NASH^[323]71–[324]73.
Targeting NADPH oxidase to reduce ROS production is thought to be more
effective in treating ROS-related vascular disease than non-enzymatic
anti-oxidants such as vitamin E^[325]74. In this study, we show that
pharmacological activation of PRDX1’s peroxidase activity with RA is
efficient in reducing H[2]O[2] levels and improving NASH, establishing
a proof of concept that boosting the peroxidase activity of PRDX1
ameliorates NASH, and indicating that PRDX1 is a promising therapeutic
target. Furthermore, our study suggests that activation of anti-oxidant
enzymes could be a feasible way to combat oxidative stress and related
human diseases including NASH, diabetes, atherosclerosis, and
cardiovascular disease.
As a common and abundant saturated FFA^[326]75, PA displays potent
lipotoxicity in NASH by inducing a variety of biological effects
including inflammation and oxidative stress^[327]35. Indeed, we
observed a significant increase of H[2]O[2] levels in HepG2 cells
shortly after PA treatment. Although numerous studies have uncovered
the molecular mechanisms underlying PA’s pro-inflammatory
effects^[328]76,[329]77, little is known about the molecular basis of
PA-induced oxidative stress. One previous study has suggested that PA
stimulates oxidative stress by causing mitochondrial dysfunction and
increasing ROS production from mitochondria in liver cells^[330]64;
however, how PA causes mitochondrial dysfunction remains unclear.
Our results demonstrate that PA directly targets and inhibits the
peroxidase PRDX1, providing a sound molecular basis underlying PA’s
lipotoxicity in stimulating oxidative stress and NASH. This regulation
system could exist in NASH with or without obesity (Supplementary
Fig. [331]15). In NASH with obesity (e.g., WD feeding), systemic
insulin resistance induces lipolysis and FFA release in adipose tissue,
and stimulates de novo lipogenesis in the liver^[332]4, which inhibits
PRDX1 peroxidase activity and increases H[2]O[2] levels in the liver.
In NASH without obesity (e.g., MCD feeding), a decrease in hepatic
PRDX1 peroxidase activity could be caused by PA that is increased
primarily in the liver, as increased FA uptake has been suggested to
contribute to MCD-induced NASH^[333]78. It is also possible that PA
inhibits other anti-oxidants to promote oxidative stress and NASH,
given that the activities of anti-oxidant enzymes such as catalase and
superoxide dismutase have been shown to be significantly reduced in
NASH patients^[334]16,[335]47. Thus, as a potential scenario,
PA-induced lipotoxicity stimulates hepatic oxidative stress and
exacerbates NASH by inhibiting the whole anti-oxidant defense system.
Our study has indicated that PRDX1 protects against NASH by scavenging
H[2]O[2], mitigating the oxidation of PTPs, and ultimately suppressing
STAT1 and STAT3 signaling. Tiganis group has elegantly shown that the
phosphorylation of distinct STAT (e.g., STAT1, STAT3 and STAT5) is
significantly increased in HFD-induced NAFL or NASH, which is ascribed
to the oxidation of PTPs (in particular TCPTP) caused by
ROS^[336]41,[337]42. They further demonstrated that STAT1 signaling
accounted for NASH with obesity^[338]42. With regard to STAT3
signaling, previous studies have suggested that it promotes NASH and
fibrosis^[339]79,[340]80. Consistent with these findings, our results
support that increased oxidization of PTPs and phosphorylation of STAT1
and STAT3 contribute to NASH. Our results also indicate that one common
driver is H[2]O[2], as H[2]O[2] treatment in HepG2 cells markedly
increased PTPs’ oxidation and STAT1 and STAT3 phosphorylation, while
PRDX1 overexpression or pharmacological activation of PRDX1 with RA
significantly reduced hepatic H[2]O[2] levels and hepatic STAT1 and
STAT3 phosphorylation. Note that Dr. Dick group has elegantly
demonstrated a redox relay between PRDX2 and STAT3, resulting in STAT3
oxidation and inactivation^[341]14,[342]81; however, whether a similar
redox relay between PRDX1 and STAT1 or STAT3 that blocks their
activities occurs remains unknown. In addition, how the redox relay
between PRDX1 and STAT1 or STAT3 gets involved in NASH pathogenesis
remains unclear, given that STAT1 activation and STAT3 activation have
been shown to promote obesity-related NASH and HCC,
respectively^[343]42.
In addition to suppressing STAT signaling, PRDX1 protects liver
mitochondrial function by mitigating lipid peroxidation in the liver
mitochondria. Using O2k system, we found that liver mitochondria
respiratory and coupling efficiency reflected by RCR or LCR was
impaired in NASH mouse models, which is in agreement with one study
carried out in humans^[344]47. Interestingly, this study also indicated
that liver mitochondrial O[2] flux increases in NAFL patients with
obesity but decreases in NASH patients^[345]47, suggesting a
compensatory action of liver mitochondria to counteract disease
progression. Consistently, this same group recently showed that liver
mitochondrial O[2] flux was significantly increased in NASH patients
with obesity, whereas this enhancement was impaired in NASH patients
with T2D^[346]82. Whether liver mitochondrial O[2] flux in NASH mouse
models shows the same pattern as that in humans remains to be further
defined.
It is worth noting that PRDX1 showed differential effects on body
weight of mice fed a WD or MCD. PRDX1 overexpression decreased body
weight of WD-fed mice by increasing their energy expenditure
(Fig.[347]4b, d), while it had no effect on body weight of MCD-fed mice
(Supplementary Fig. [348]4d). We suppose that PRDX1 may get involved in
weight control dependent of some adipokines from adipose tissue, given
that WD feeding promotes considerable fat and weight gain, whereas MCD
feeding causes substantial fat and weight loss. Nevertheless, more
studies in future are needed to understand the molecular mechanisms
behind differential weight control by PRDX1.
In summary, we demonstrate in the study that PA promotes hepatic
oxidative stress and NASH by binding PRDX1 and inhibiting its
peroxidase activity, and PRDX1 protects against NASH through its
peroxidase activity. Hence, activation of PRDX1’s peroxidase activity
is a promising way to treat NASH.
Methods
Animals
All animal studies were conducted in strict accordance with the
standards of animal welfare and institutional guidelines for the humane
treatment of animals, and were approved by the Animal Care and Use
Committee (ACUC) at Chu Hsien-I Memorial Hospital & Tianjin Institute
of Endocrinology, Tianjin Medical University (DXBYY-IACUC-2020036). The
mice were euthanized with CO[2] or isoflurane inhalation, followed by
cervical dislocation at the end of the experiment. Male mice were used
for phenotypic and mechanistic analyses throughout the study.
All mice (C57BL/6 J background) were housed at a facility with
controlled temperature (22 °C), humidity and a 12/12 hr light/dark
cycle, and had ad libitum access to food and water. C57BL/6 WT mice
were purchased from GemPharmatech (Nanjing, China). Alb-Cre (Stock #:
003574) mouse strain was obtained from the Jackson Laboratory (Bar
harbor, ME)^[349]39. PRDX1 KO (Prdx1^-/-) and overexpression
(Prdx1^OE/OE) mice were generated by GemPharmatech (Nanjing, China)
using CRISPR–cas9 approach. To generate Prdx1^-/- mice, single guide
RNAs (sgRNAs) targeting the exons 2–5 of Prdx1 were designed. To
generate Prdx1^OE/OE mice, a donor vector harboring the mouse Prdx1
coding sequence (CDS) in fusion with an HA tag was inserted into the
ubiquitous H11 site. Loxp-flanked Prdx1 (Prdx1^fl/fl) mouse strain was
generated by GemPharmatech (Nanjing, China). To generate Prdx1^fl/fl
mouse strain, two loxp sequences were inserted and flanked the exon 3
(E3) of Prdx1 gene via CRISPR-Cas9 system. PRDX1^Cys52Ser mice were
originally generated and maintained in our laboratory^[350]54.
Different NAFLD mouse models were induced by high-fat diet (HFD,
Research Diets, D12492, 60 kcal% Fat), methionine and choline-deficient
diet (MCD, MolDietes, M0421), western diet (WD, Research Diets,
D09100310, 40 kcal% Fat, 20 kcal% Fructose and 2% Cholesterol),
choline-deficient, amino acid-defined, HFD (CDAHFD, Research Diets,
A06071302), or customized CDAHFD containing different amounts of PA
(MolDietes) (Supplementary Table [351]3) feeding for a time as
specified in the text.
Cell culture and plasmid construction
HepG2 (ATCC:HB-8065, Manassas, VA, USA) and HEK293T cells
(ATCC:CRL-3216, Manassas, VA, USA) were grown at 37 °C in Dulbecco’s
Modified Eagle Medium (DMEM, GIBCO) supplemented with 10% fetal bovine
serum (FBS, GIBCO), 100 IU/ml penicillin and 100 mg/ml streptomycin.
To examine the effects of N-Acetyl-L-cysteine (NAC, Sigma) on the
phosphorylation of STAT1 and STAT3 stimulated by IL-6 (206-IL-010, R&D
Systems) or IFN-γ (ab9659, Abcam), HepG2 cells were incubated with
serum-free medium for 30 min, and pretreated with NAC (5 mM) for
30 min, followed by treatment with IL-6 (10 ng/ml) or IFN-γ (10 ng/ml)
for another 30 min.
To assess the effect of RA on LPS-stimulated PRDX hyperoxidation
(SO2/SO3), HepG2 cells were incubated with serum-free medium for
30 min, and then pretreated with RA (1 μM) for 3 hr, followed by veh or
LPS (100 ng/ml) treatment for another 3 hr.
To assess the anti-inflammatory effect of RA, primary hepatocytes were
isolated from 6-week-old C57BL/6 J mice as previously
described^[352]54. The primary hepatocytes were cultured in RPMI 1640
medium containing 10% FBS for 24 hr after isolation, and then replaced
with serum-free medium followed by treatment with veh, LPS (100 ng/ml)
or RA (1 μM) for 6 hr.
Mutation of PRDX1 Cys52 to Ser52 (PRDX1^Cys52Ser) was achieved by
PCR-directed mutagenesis as previously described^[353]83. In brief, two
rounds of PCR were conducted through different pairs of primers to
mutate Cys52 of PRDX1 to the Ser residue. To generate the plasmids
expressing PRDX1, PRDX1^Cys52Ser or Ppp1ca, the mouse coding sequence
(CDS) was inserted in pcDNA3.1-3xFLAG or pcDNA3.1-3xHA empty vectors
(YouBio, China) with epitope fusion at the N-terminus. All DNA
constructs were sequenced, and transfected in 293 T cells to confirm
their protein expression before IP-MS or Co-IP analyses. All PCR
primers used in the study are listed in the Supplementary Table [354]4.
Measurement of PRDX peroxidase activity
Global PRDX peroxidase activity in the liver or HepG2 cell lysate
(freshly prepared) was measured with a classic Trx-TrxR-NADPH coupled
assay as previously described^[355]28,[356]29 with some modifications.
In brief, 200 μM NADPH (Sigma), 3 μM Trx1, and 1.5 μM TrxR1 were added
in 50 mM HEPES-NaOH buffer (pH 7.0) containing 100 ug of total protein.
Note that both yeast Trx1 and yeast TrxR 1 were purified by our own
laboratory. 50 μM H[2]O[2] was added to initiate the reaction at 30 °C,
followed by detection of absorbance at 340 nm (A[340]) every 20 s for
15 min assay duration. The background activity was simultaneously
assessed without Trx and TrxR, but only with H[2]O[2] and NADPH. To
calculate the initial NADPH consumption rate (initial rate)
(A[340]/min/protein (g)) in the first 5 min, a smooth curve was drawn
through A[340] readings, and the initial rate was calculated by
performing a simple linear regression (GraphPad Prism 9). The global
PRDX peroxidase activity was calculated by subtracting the background
activity (initial rate) from total activity (initial rate).
Recombinant WT PRDX1 peroxidase activity assay was conducted as
previously described^[357]84,[358]85. To measure the effect of sodium
palmitate on recombinant WT PRDX1’s peroxidase activity, recombinant WT
PRDX1 (400 nM) was incubated with sodium palmitate for 30 min before
addition of a mixture buffer containing 3 μM Trx, 1.5 μM TrxR, and
200 μM NADPH. NADPH reduction was monitored via A[340] right after the
addition of 100 μM H[2]O[2] for 60 min. The quantitative analysis was
performed as we previously described^[359]85. Briefly, the slope of
each assay well was calculated from linear part of the curve. The slope
of assay wells without PA and PRDX1 was considered as full inhibition
(S[100%]) and the slope of assay wells containing PRDX1 but no PA was
considered as no inhibition (S[0%]). For PA at each concentration, the
% inhibition was calculated according to the following equation: %
inhibition = 100 –100 x (S[PA] − S[100%]) / (S[0%] − S[100%]). The data
were processed by GraphPad Prism 9.
HKPerox-Red staining in liver sections and cells
Liver samples were frozen and embedded in OCT compound (SAKURA).
Cryosections were prepared at the thickness of 8 μm for HKPerox-Red
staining. Frozen sections were incubated with 5 µM HKPerox-Red in PBS
(0.1% DMF(Macklin), 100 mM CCl3CN (Macklin)) for 10 min at room
temperature, followed by staining with DAPI (1 μg/ml) (Sigma) for
5 min. HKPerox-Red staining was performed in live cells as previously
described^[360]31. In brief, HepG2 cells were treated with PA at
different concentrations (250 µm or 500 µm) for 3 hrs, followed by
incubation with HKPerox-Red (10 mM) in Hank’s Buffer (0.1% DMF, 100 mM
CCl3CN) at 37 °C for 30 min. Images were captured with an Olympus
fluorescence microscope. To quantify the fluorescence intensity of
images, Image J was used to convert RGB to 8-bit format, adjust
fluorescence threshold, and measure the integrated density. Finally,
the data were processed by GraphPad Prism 9.
Measurement of malondialdehyde (MDA) levels
To quantify the extents of lipid peroxidation in the liver, isolated
mitochondria, or HepG2 cells, a commercial MDA detection kit (Beyotime)
was applied to measure MDA concentrations according to the
manufacturer’s instructions. Briefly, the mixture of samples with MDA
detection buffer was heated at 100 °C for 10 min, and then centrifuged
at 1000 x g for 10 min at room temperature. The supernatant contained
MDA-TBA adduct formed after a chemical reaction between MDA and
thiobarbituric acid (TBA). MDA-TBA adduct has a maximal absorbance at
535 nm, which can be monitored by a fluorometer (BioTek). MDA
concentration was finally normalized to protein concentration (mg/ml).
Physiological measurements
Measurements of weekly body weight and daily food intake were carried
out as previously described^[361]86. Energy expenditure (kcal) and
locomotion activity were monitored using Promethion High-Definition
Multiplexed Respirometry System (Sable Systems, North Las Vegas, NV,
USA).
Serum AST and ALT levels were analyzed with an automatic blood
biochemical analyzer (AU5800, Beckman Coulter).
Histological analyses
Liver samples were fixed with 4% paraformaldehyde, embedded in paraffin
and prepared for H&E, Sirius Red, or immunohistochemical staining.
Staining of H&E and Sirius Red was performed with individual kits from
Solarbio and Leigen companies according to the manufacturer’s
instructions.
Immunohistochemical staining was performed as previously
described^[362]87. In brief, a rabbit polyclonal antibody against α-SMA
(1:1000 dilution) (14395-1-AP, Proteintech) was incubated with the
paraffin-fixed liver sections at 4°C overnight, followed by the
incubation with a polymer-HRP anti-rabbit secondary antibody and
detected with DAB (3,3’-diaminobenzidine) stain obtained from
Proteintech (PK10006). An Olympus fluorescence microscope with 20x or
40x objective lens was used to capture images.
Liver samples were frozen and embedded in OCT compound (SAKURA).
Cryosections were prepared at the thickness of 8 μm for Oil Red O
staining as previously described^[363]88.
Western blotting
Western blotting was performed as previously described^[364]86.
Briefly, tissues or cultured cells were homogenized with 1 x lysis
buffer containing 1% deoxycholic acid, 10 mM Na[4]P[2]O[7], 1% Triton
100, 100 mM NaCl, 5 mM EDTA, 50 mM Tris-HCl, and 0.1% SDS. Protein
concentrations were determined through BCA protein assay (23228, Thermo
Fisher Scientific, Rockford, IL, USA). In general, 20–40 μg of protein
was loaded in SDS-PAGE and transferred to PVDF membranes. After
blocking with 5% nonfat milk in TBST for 1 hr, PVDF membranes were
incubated with the primary antibodies overnight at 4 °C, followed by
secondary antibodies for 1 hr at room temperature (Fig. [365]5c–g). ECL
detection systems were applied to develop signals.
To detect in vitro PRDX1 hyperoxidation (SO2/SO3), recombinant WT PRDX1
(100 ng) was incubated with RA at different concentrations for 30 min
before addition of a mixture buffer containing 1.5 μM Trx, 0.8 μM TrxR,
and 200 μM NADPH. The reaction was initiated with 5 μM H[2]O[2]. After
8 min, the reaction was quenched by 2 x sample buffer followed by
SDS-PAGE and western blotting.
Antibodies
Antibodies used in this study include PRDX1 rabbit monoclonal antibody
(Cell Signaling, 8499), PRDX2 rabbit polyclonal antibody (Thermo
Fisher, PA5-86019), PRDX3 rabbit polyclonal antibody (Proteintech,
10664-1-AP), PRDX4 rabbit polyclonal antibody (Proteintech,
10703-1-AP), PRDX5 rabbit polyclonal antibody (Proteintech,
17724-1-AP), PRDX6 rabbit polyclonal antibody (Proteintech,
13585-1-AP), Smooth muscle actin (α-SMA) rabbit polyclonal antibody
(Proteintech, 14395-1-AP), GAPDH monoclonal antibody (Proteintech,
60004-1-Ig), Oxidized PTP active site mouse monoclonal antibody (R&D
Systems, MAB2844), β-Actin mouse monoclonal antibody (Sigma-Aldrich,
A5441), Peroxiredoxin-SO3 rabbit polyclonal antibody (Abcam, ab16830),
Phospho-STAT1 (Tyr701) (58D6) rabbit antibody (Cell Signaling, 9167),
STAT1 rabbit polyclonal antibody (Cell Signaling, 9172), Phospho-STAT3
(Tyr705) (D3A7) rabbit antibody (Cell Signaling, 9145), STAT3 (D3Z2G)
rabbit antibody (Cell Signaling, 12640), HA-tag (C29F4) rabbit antibody
(Cell Signaling, 3724), FLAG tag rabbit antibody (Proteintech,
80010-1-RR), Phospho-STAT1 (Y701) rabbit antibody (ABclonal, AP0054),
F4/80 rabbit antibody (Cell Signaling, 70076), Phospho-WWTR1(Ser89)
rabbit antibody (Invitrogen, PA5-105066), PPP1ca mouse antibody
(Proteintech,67070-lg), Phospho-PPP1ca (Thr320) rabbit antibody
(Proteintech, 29874-1-AP), YAP/TAZ(D24E4) rabbit antibody (Cell
Signaling,8418), Goat Anti-Rabbit IgG Antibody, (H + L) HRP conjugate
(Millipore, AP187P), and Goat Anti-Mouse IgG Antibody, HRP conjugate
(Millipore, AP181P).
Measurement of intracellular ROS
According to the manufacturer’s instructions, intracellular ROS in
HepG2 cells were measured by detecting the fluorescent intensity of
dichlorofluorescin (DCF), an oxidized product from non-fluorescent
compound 2’,7’-dichlorodihydrofluorescein diacetate (H2DCFDA)
(ThermoFisher). Briefly, HepG2 cells were starved in serum-free Hank’s
buffer for 30 min, followed by stimulation with PA (250 μM) for another
30 min. To evaluate the anti-oxidative effect of RA, HepG2 cells were
starved in serum-free Hank’s buffer and simultaneously pretreated with
RA (1 μM) for 30 min, followed by stimulation with IL-6 (10 ng/ml) for
another 30 min. Afterward, cells were first incubated with 5 μM H2DCFDA
at 37 °C for 30 min in the darkness and then detected at 485 nm
(excitation) and 520 nm (emission) by a fluorometer (BioTek).
Surface plasmon resonance (SPR)
The SPR binding assay was performed on a Biacore T200 instrument.
Recombinant wild-type PRDX1 at 200 μg/ml in 10 mM sodium acetate (pH =
4.5) was coupled with the CM5 chip (GE Healthcare). After
immobilization, the system was equilibrated for 1 hr. RA was injected
and flowed through the chip at a flow rate of 20 μl/ml in assay buffer
with 0.05% Tween-20. As for sodium palmitate, SPR assay was performed
in PBS with 0.01% NP-40. Each injection was associated with the sensor
chip for 120 s and dissociated for 180 s. All data were processed using
the Biacore T200 Evaluation software (version 1.0).
Cellular thermal shift assay
The cellular thermal shift was performed in HepG2 cells as previously
described^[366]36. In brief, cells were treated with PA (Sigma) that
was prepared in 10% BSA (fatty acid free, Sigma) or 10% BSA as a
control for 1 hr, and then harvested and suspended in 1x PBS containing
protease inhibitors (cocktail, Roche). Cell suspensions were divided
into a number of aliquots (50 μl) that were heated at different
temperatures ranging from 55.5 °C to 64.0 °C for 3 min, followed by
cooling at room temperature for 3 min. Heated cell suspensions were
freeze-thawed with liquid nitrogen for 3 times before they were
centrifuged at 20,000 x g at 4°C for 30 min. The supernatants were
collected for SDS-PAGE and western blotting analyses.
Quantitative PCR (qPCR)
qPCR was performed and quantified as previously described^[367]86. In
brief, mouse tissues or cultured cells were homogenized in TRIzol
(Invitrogen) and total RNA was extracted. In general, 1 µg RNA in total
was used for reverse transcription with random primers, followed by
quantitative PCR with QuantStudio 3 Real-Time PCR System (Applied
Biosystems). The relative expression of target genes was calculated
based on 2^-ΔΔCt method with 36b4 as the reference gene. Mouse primers
used in this study were summarized in Supplementary Table [368]2.
IPGTT and IPITT
Intraperitoneal glucose tolerance test (IPGTT) and intraperitoneal
insulin tolerance test (IPITT) were performed as previously
described^[369]86. For IPGTT, mice were fasted overnight (16 hr) and
the fasting glucose levels were measured right before mice were
intraperitoneally injected with glucose (1.0 g/kg body weight). After
glucose injection, blood glucose levels were measured every 30 min
until 2 hr post injection. For IPITT, animals were fasted for 6 hr. The
fasting glucose levels were measured before an intraperitoneal
injection of insulin (1.5 U/kg body weight). Blood glucose levels were
measured every 30 min until 2 hr post injection of insulin. A portable
glucometer (OneTouch Ultra) was used to measure blood glucose.
Measurement of hydrogen peroxide (H[2]O[2])
To measure the concentration of H[2]O[2] in the liver or HepG2 cells,
we employed a hydrogen peroxide assay kit (ab102500, abcam).
Measurement was performed according to the manufacturer’s instructions.
In brief, liver or cell samples were collected fresh, washed in cold
PBS, and lysed in the assay buffer. Supernatant was then subject to
deproteinization to remove proteins using a commercial deproteinizing
sample preparation kit (ab204708, abcam). Following protein removal,
the supernatant was used for fluorometric assay at the excitation
wavelength of 535 nm by a fluorometer (BioTek). H[2]O[2] concentration
was calculated according to the manual provided.
Generation of PRDX1 knockout HepG2 cells
To generate PRDX1 knockout HepG2 cells, we employed CRISPR-Cas9
approach. Lentivirus (pHBLV-U6-hPrdx1-gRNA-EF1-CAS9-PURO) expressing
Cas9 and gRNA for human PRDX1 was packaged in 293 T cells by HANBIO
(Shanghai, China) according to the standard procedure. HepG2 cells were
infected with the packaged lentivirus and PRDX1 knockout HepG2 cells
were screened out through puromycin treatment. The knockout efficiency
was confirmed by western blotting. The gRNA sequence used in this
study: CCTGAGCAATGGTGCGCTTC (5’-3’).
Transcriptome analysis
Liver samples were collected from mice and stored in RNAlater (Ambion)
overnight. Total RNA was isolated and purified for transcriptome
analysis. In brief, RNA was isolated using TRIzol reagent and RNA
quality was evaluated with Bioanalyzer 2100 (Agilent, CA, USA). With
high-quality RNA, cDNA library was created and then the 2x150bp
paired-end sequencing was performed on an Illumina Novaseq 6000 (LC-Bio
Technology Co., Hangzhou, China).
For bioinformatics analysis, fastp software
([370]https://github.com/OpenGene/fastp) was employed to remove
unnecessary reads and verify sequence quality. [371]HISAT2 was applied
to map reads to the reference genome of mus_musculus/Ensembl/v101 and
generate bam files. StringTie
([372]https://ccb.jhu.edu/software/stringtie) was used to assemble and
quantify the mapped beads of each sample with default parameters.
Gffcompare ([373]https://github.com/gpertea/gffcompare/) was used to
merge all transcriptomes to reconstruct a comprehensive transcriptome,
followed by estimating the expression levels of all transcripts using
StringTie. R package edgeR
([374]https://bioconductor.org/packages/release/bioc/html/edgeR.html)
was used to select differentially expressed genes (DEGs). DAVID
software ([375]https://david.ncifcrf.gov/) and GSEA4.1.0 software
([376]http://www.gsea-msigdb.org/gsea/index.jsp) was applied for Kyoto
Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analysis
and gene-set enrichment analysis (GSEA), respectively.
Isolation of liver mitochondria
Isolation of fresh liver mitochondria for Oxygraph-2k (O2k) study was
performed according to a previous study with some
modifications^[377]89. In brief, approximately 500 mg liver tissue was
collected in 2–3 ml of pre-cold mitochondria isolation buffer (225 mM
mannitol (Sigma), 75 mM sucrose (Sigma), 0.2 mM EDTA (Solarbio)),
followed by homogenization for 10–12 times using a Teflon-glass
homogenizer. The homogenates were subject to centrifugation at 1000 x g
for 10 min at 4 °C and the supernatants were collected for a second
centrifugation at 6200 x g for 10 min at 4 °C. The resultant
mitochondrial fraction was suspended in 1 ml of pre-cold Mir05
mitochondrial respiration medium for later O2k analyses.
The procedure for isolating liver mitochondria for MDA measurement was
similar to that described above except the followings: after
centrifugation at 6200 x g for 10 min, the pellet was lysed with 0.5 ml
lysis buffer (1% Triton X-100, 0.1% SDS), followed by the
centrifugation at 13,800 x g for 10 min at 4 °C. The supernatant was
collected for measurement of MDA and protein concentration.
Assessment of mitochondrial citrate synthase activity
Mitochondrial citrate synthase activity (CSA) was measured according to
a protocol from Oroboros
([378]https://wiki.oroboros.at/index.php/MiPNet17.04_CitrateSynthase).
Briefly, mitochondrial suspension was mixed with buffer (1 M Tris-HCl,
1 mM EDTA, 0.25% Triton X-100, 0.31 mM acetyl-CoA, 0.1 mM
5,5’-dithiobis (2-nitrobenzoic acid) and 0.1 M triethanolamine) and the
mixture was then added with 0.5 mM oxalacetate to initiate the
reaction. A spectrophotometer was applied to record the absorbance at
412 nm at 37 °C every 20 s over a 10-min period. CSA was calculated
based on the following equation:
[MATH:
v=rAl⋅ε
B⋅vB⋅VcuvetteVsample⋅ρ :MATH]
V: specific activity of the enzyme (IU/mg protein); r[A]: rate of
absorbance change (dA/dt); ε[B]: extinction coefficient of B (TNB) at
412 nm and pH 8.1(13.6 mM^-1.cm^-1); V[B]: stoichiometric number of B
(TNB in the reaction) (V[B] = 1); Vcuvette: volume of solution in the
cuvette; Vsample: volume of sample added to cuvette; ρ: mass
concentration or density of biological material in the sample (protein
concentration: mg. ml^-1).
Measurement of liver mitochondrial O[2] flux
We used Oxygraph-2k (O2k) (Oroboros, Austria) to assess liver
mitochondrial respiration. Liver mitochondrial were isolated fresh as
described above and used for oxygen (O[2]) flux measurement in Oroboros
chambers containing respiration buffer (MiR05). Approximately 100 μg of
total mitochondria were loaded in one chamber for every measurement.
The high-resolution respirometry (HRR) protocol for measuring O[2] flux
in isolated mitochondria: malate (Sigma), glutamate (Sigma), ADP
(Sigma), succinate (Sigma), cytochrome c (Sigma), CCCP (Sigma). O[2]
flux was monitored at different states after adding substrates or
inhibitors. Hepatic mitochondrial content was determined according to
CSA. Mitochondrial O[2] flux was presented as the value that was
normalized to the relevant CSA.
Measurement of liver mitochondrial respiratory and coupling efficiency
As defined in one previous study^[379]47, respiratory control ratio
(RCR) and leak control ratio (LCR) was calculated as the ratio of state
3 over state o and the ratio of state o over state u, respectively. We
employed O2k (Oroboros, Austria) to evaluate RCR and LCR in fresh liver
mitochondria via the following protocol: malate, glutamate, ADP,
succinate, oligomycin (Sigma), CCCP. Oxygen flux at different states:
ADP-stimulated coupled respiration (state 3); respiration after adding
oligomycin (state o), and maximal uncoupled respiration after addition
of uncoupling CCCP. Approximately 100 μg of total mitochondria were
loaded in one chamber for every measurement.
Compound library screening via protein thermal shift (PTS)
Polyphenolic natural compound library was purchased from TargetMol.
Before screening procedures, recombinant WT PRDX1 was reduced by TCEP
(Tris(2-carboxyethyl)phosphine). After reduction, PRDX1 was desalted
into assay buffer (20 mM Hepes 7.0, 150 mM NaCl). Working concentration
of recombinant WT PRDX1 and compounds were 10 μM and 800 μM
respectively for first and second round screens. 5×SYPRO orange dye was
mixed with PRDX1 and then compounds were added before melting
temperature detection by QuantStudio™ 6 Flex Real-time PCR system
(Applied Biosystems). The fluorescence signal was collected with
gradient elevation of heating temperature from 25 °C to 95 °C for
25 min. Delta melting temperature (ΔT[m]B and ΔT[m]D) were calculated
using assay wells containing PRDX1 without compounds as a reference
with Protein Thermal Shift™ Software (version 1.2). After the first
round screen, 16 compounds stood out as hits and 15 (one compound out
of stock) were purchased from TargetMol or CSNPharm. All 15 hits were
dissolved in DMSO at 20 mM and rescreened using the same procedure as
the first round screen. Data were analyzed by GraphPad Prism 9.
Measurement of activation of PRDX1’s peroxidase activity
To measure the effects of candidate compounds from library screening on
PRDX1’s peroxidase activity, compounds were incubated with PRDX1
(400 nM) for 0.5 hr and then mixed with pre-reaction mixture containing
1.5 μM Trx, 0.8 μM TrxR and 200 μM NADPH. The reaction was initiated
with 200 μM H[2]O[2] at room temperature, followed by addition of
ROSGreen^TM H[2]O[2] probe^[380]90–[381]92 (MX5202, MKBio, China) in
each well for H[2]O[2] quantification. Activation of PRDX1’s peroxidase
activity by each compound was calculated from fluorescence intensity
detected by ROSGreen^TM H[2]O[2] probe. To obtain the real
fluorescence, the fluorescence intensity of assay wells containing
PRDX1 and RA minus the fluorescence intensity of RA only. The delta
fluorescence intensity (ΔF) was calculated as fluorescence intensity of
assay wells containing pre-reaction mixture and H[2]O[2] without PRDX1
and compounds (F[blank]) minus the fluorescence intensity of assay
wells containing pre-reaction mixture, H[2]O[2] and PRDX1 incubation
with or without compounds (F[PRDX1+compound], or F[PRDX1-compound]),
which reflects H[2]O[2] consumption of each well
(ΔF=F[blank]–F[PRDX1+compound,] or F[blank]–F[PRDX1-compound]). To
quantify the relative H[2]O[2] consumption, ΔF of PRDX1 with compound
was divided by that of PRDX1 without compound (fold of H[2]O[2]
consumption =ΔF (F[blank] – F[PRDX1+compound]) / ΔF (F[blank] –
F[PRDX1-compound]). Finally, the compound’s half-maximum concentration
for activating peroxidase activity was calculated by GraphPad Prism 9.
Protein expression and purification
cDNAs of human WT PRDX1 protein (aa 1-199) and its truncation mutant
(C52SC83S, aa 1-175) for crystallization were inserted into vector
pet28a (+) with a N terminal 6× his tag and a TEV protease cleavage
site. Then the constructed plasmids were transformed into E.coli strain
BL21 (DE3). The expression of PRDX1 was induced with 400 mM IPTG at 16
°C overnight. After ultrasonication in buffer A (50 mM Tris7.0, 200 mM
NaCl, 20 mM imidazole, 10 mM β-mercaptoethanol), the clear lysate was
loaded into 5 ml his trap HP column (Cytiva) and eluted with 700 mM
Imidazole in buffer A by AKTA Pure. The protein was desalted in assay
buffer (20 mM Hepes 7.0, 150 mM NaCl) and stored at -80 °C with 5%
glycerol. For crystallization, his tag was removed by TEV protease and
further purified by size exclusive chromatography Superdex 75 Increase
10/300 GL (Cytiva). Protein quality was assessed with SDS-PAGE.
Crystallization, data collection and structure determination
PRDX1^C52SC83S (aa1-175) at 5 mg/ml was crystallized in buffer (20 mM
Tris 8.5, 100 mM NaCl, 1 mM TCEP) by sitting-drop method at 16 °C using
a reservoir solution of 10% v/v Tacsimate pH 7.5, 0.1 M MES pH 6.5, and
25% PEG4000. Ligand free crystal was soaked in reservoir solution with
the addition of 2 mM RA overnight. For data collection, the crystal was
protected by cryo-protectant solution containing 25% glycerol and then
flash frozen in liquid nitrogen.
X-ray diffraction data were collected at the BL19U1 beamline of
National Facility for Protein Science in Shanghai (NFPS) at Shanghai
Synchrotron Radiation Facility. Full 360° diffraction data were
collected with a detector distance of 350 mm. The data were processed
using XDS for integration and CCP4 for scale. The structure of
PRDX1^C52SC83S (aa1-175) was solved by molecular replacement using
PHENIX (version 1.19.2), with the dimer of PRDX1 (PDB code: 9B7A) as
the search template. RA was prepared using eLBOW module and placed
using LigandFit module in PHENIX. The initial refinement was carried
out in PHENIX and then checked manually in Coot (version 0.9.4). Data
collection and refinement statistics are shown in Supplementary
Table [382]1.
Administration of RA in mice
To evaluate the therapeutic potential of RA (CSNpharm) in improving
NASH, RA was prepared fresh in saline and injected (30 mg/kg) once
daily intraperitoneally in mice that concurrently were fed a specific
diet to induce NASH.
Analysis of lipid peroxidation by liquid chromatography (LC) mass
spectrometry (MS)
Lipid peroxidation analysis was performed as described^[383]93.
Briefly, PE (D16:1) and PC (D14:1) were used as internal standards in
each sample, and lipids were extracted using the Folch method. Next,
0.005% BHT in ice-cold chloroform/methanol (v/v = 2:1) was added to
samples, vortexed and incubated on ice for 15 min. After
centrifugation, the lower organic layers were collected in a new tube
and dried under N[2] flux. The dried samples were resuspended in 60 µL
of 100% LC solvent B, followed by being aliquoted and transferred to a
new autosampler vial for analysis.
Mass spectrometric analysis was performed in the multiple-reaction
monitoring (MRM) of specific precursor–product ion m/z transitions upon
collision-induced dissociation. The precursor negative ions monitored
were the molecular ions [M − H]− for PE, and the acetate adducts
[M + CH3COO]− for PC. Meanwhile, identity was verified by monitoring.
The positive molecular ions [M + H]+ for both PC and PE were monitored
simultaneously using polarity switching.
Immunoprecipitation (IP)-mass spectrometry (MS) and Co-IP
To identify proteins differentially interacting with WT PRDX1 and
PRDX1^Cys52Ser mutant, pcDNA3.1-3xFlag-PRDX1 and
pcDNA3.1-3xFlag-PRDX1^Cys52Ser expression constructs were transfected
in HEK293T cells for 48 hr. Afterward, cells were harvested and lysed
in 1% NP-40 lysis buffer (20 mM Tris-HCl, 0.5% NP40, 150 mM NaCl, 1 mM
EDTA with complete protease inhibitor cocktail (Roche) and phosphatase
inhibitor (PhosSTOP, Roche)). Cell lysates were used for
immunoprecipitation (IP). Briefly, 4 mg of total protein was incubated
with 50 μL of Anti-Flag M2 affinity gel (Sigma, A2220) at 4°C for
overnight, followed by washing with 1x lysis buffer six times. After
the final wash, the precipitated protein samples were denatured in
2xSDS sample buffer at 100 °C for 10 min, followed by SDS-PAGE and
coomassie blue staining.
The subsequent proteome analysis was carried out by Jingjie
Biotechnology Co. (Hangzhou, China). For in-gel tryptic digestion,
coomassie blue-stained gel bands were cut and destained in 50 mM
NH[4]HCO[3] in 50% acetonitrile (v/v) until clear. The gel pieces were
dehydrated with 100% acetonitrile for 5 min, and rehydrated in 10 mM
DTT at 37 °C for 60 min. After two additional rounds of dehydration and
rehydration, the gel pieces were digested with trypsin (10 ng/μl) at
37°C overnight. Peptides were extracted, and then dried to completion
and resuspended in 2% acetonitrile/0.1% formic acid.
For mass spectrometry (MS) analysis, the tryptic peptides were
separated by EASY-nLC 1200 UPLC system and analyzed through Orbitrap
Exploris 480 MS. The MS data were processed using Proteome Discoverer
2.4.
To validate the interaction between PPP1ca and PRDX1 or PRDX1^Cys52Ser,
plasmids expressing 3xFlag tagged protein or 3xHA tagged protein were
co-transfected in 293 T cells for Co-IP analysis. 48 hr post
transfection, cells were harvested for IP using Anti-Flag M2 affinity
gel or normal mouse IgG magnetic beads as described above. Finally, the
precipitated protein samples were analyzed by WB to investigate protein
interaction.
Statistical analysis
All results are presented as means ± SEM. ImageJ was used to quantify
protein levels from western blotting images, or fluorescence intensity
from microscopic images. Statistical analyses were performed with
unpaired and two-tailed Student’s t test (Excel 2011), or one-way and
two-way ANOVA followed by the Bonferroni test for multiple comparisons
(GraphPad Prism 9). p < 0.05 was considered statistically significant.
GraphPad Prism 9 and Adobe Illustrator CS 2020 were used to generate
and prepare all figures.
Reporting summary
Further information on research design is available in the [384]Nature
Portfolio Reporting Summary linked to this article.
Supplementary information
[385]Supplementary Information^ (17.2MB, pdf)
[386]Reporting Summary^ (2MB, pdf)
[387]Transparent Peer Review file^ (3.5MB, pdf)
Source data
[388]Source Data^ (58.5MB, xlsx)
Acknowledgements