Information

What is the functional significance of the difference in cardiolipin/cholesterol ratio in different membranes?


I have read somewhere that the plasma membrane has little cardiolipin but excess cholesterol whereas the inner mitochondrial membrane is rich in cardiolipin and has little cholesterol.I just wanted to know why there is this difference.


According to wikipedia, cardiolipin is found in two places, the inner mitochondrial membrane, and bacterial membranes. Given the endosymbiotic origin of mitochondria, it makes sense that they would retain some remnant of their bacterial ancestry. But more importantly, cardiolipin plays a role in the enzymatic functions of mitchondrial membranes. Cardiolipin forms a bicyclic resonance structure that allows it to trap a proton, which helps oxidative phosphorylation.

Because this structure also responds to pH and divalent cations, it can change it's conformation and aggregation behavior as cellular conditions change.

Cardiolipin also plays a role in holding enzyme complex quaternary structures together. Mitochondrial Complex III requires 1 cardiolipin, Complex IV requires 2 cardiolipin, and Complex V binds 4 cardiolipins.

Cardiolipin also plays a role in apoptosis, where it is oxidized and undergoes conformational changes that cause it to move from inner to outer mitochondrial membranes and generate pores that cytochrome C can escape through and trigger apoptosis.

Cholesterol can be easily oxidized into oxysterols, which are thought to perform a variety of functions, but are not fully understood. If cholesterol levels in the mitochondria were high, I think it would be oxidized and cause trouble. http://en.wikipedia.org/wiki/Oxysterol


Phospholipid and cholesterol content of ventricular tissue from the cardiomyopathic Syrian hamster ☆

A relationship between the development of hereditary cardiomyopathy in the Syrian hamster (BIO 14.6) and changes in ventricular phospholipid and cholesterol content was investigated. In an attempt to distinguish purely age-related changes in lipids, young (30 day) and old (230 day) cardiomyopathic, and normal hamsters (RB) were examined. The old cardiomyopathic hamster ventricles contained more cholesterol on the basis of dry weight and non-connective tissue protein nitrogen than did normal ventricles at the same age. The phospholipid phosphorus content was lower in ventricular muscle of young myopathic hamsters relative to age-matched normals whereas there was no significant difference in older animals. Despite the loss of function of the old myopathic hearts, phospholipid composition was not markedly altered. The old myopathic ventricular muscle contained 12% less phosphatidyl ethanolamine and 14% more phosphatidyl inositol than did the old normal tissue. Cardiolipin, an indicator of mitochodrial content was unchanged.


Genetic Regulation of Intestinal Lipid Transport and Metabolism

Acyl-Coenzyme A Cholesterol Acyltransferase 2 and the Regulation of Intestinal Cholesterol Absorption

Cellular cholesterol esterification is accomplished by two enzymes: ACAT1, which is widely distributed, but expressed at low levels in the liver and intestine and ACAT2, which is the enzyme responsible for cholesterol esterification in these tissues ( 69 ). To examine the role of ACAT2 in regulating intestinal cholesterol absorption, Repa and colleagues ( 221 ) generated ACAT2 −/− mice and demonstrated reduced (but not eliminated) cholesterol absorption, particularly in the setting of increased dietary cholesterol intake. In addition, the reduction in NPC1L1 mRNA seen in wild-type animals after cholesterol feeding was amplified in ACAT2 —- mice, suggesting that the accumulation of free intracellular intestinal cholesterol further down-regulated NPC1L1 expression beyond the effects noted in wild-type mice. In addition, these workers demonstrated an increase in ABCG5/G8 mRNA abundance in chow-fed ACAT2 −/− mice, but no further increase after cholesterol supplementation, suggesting that the regulation of ABCG5/G8 expression in response to sterol supplementation involves ACAT2. A striking increase was noted in the expression of the basolateral efflux transporter ABCA1 in chow-fed ACAT2 −/− mice, and a further increase in ABCA1 mRNA abundance was noted in ACAT2 −/− mice fed a high-cholesterol diet. These data suggest that free cholesterol accumulation in enterocytes in response to ACAT2 deficiency leads to up-regulation of basolateral cholesterol efflux mechanisms, although the form of the lipoprotein particles associated with this augmented cholesterol secretion have yet to be identified. However, based on current information, the most likely candidate would be HDL particles formed by the actions of ABCA1 and ApoA–I (see Fig. 67-3 ).


The Triglyceride/HDL Cholesterol Ratio. What Is Ideal?

The TG/HDL-C ratio can easily be calculated from the standard lipid profile. Just divide your TG by your HDL-C.

However, when looking at the ideal ratio, you have to check if your lipid values are provided in mg/dl like in the US or mmol/L like in Australia, Canada, and most European countries.

If lipid values are expressed as mg/dl (like in the US)

TG/HDL-C ratio less than 2 is ideal

TG/HDL-C ratio above 4 is too high

TG/HDL-C ratio above 6 is much too high

If you are using mmol/L (most countries except the U.S.) you have to multiply this ratio by 0.4366 to attain the correct reference values. You can also multiply your ratio by 2.3 and use the reference values above.

If lipid values are expressed as mmol/L (like in Australia, Canada, and Europe)

TG/HDL-C ratio less than 0.87 is ideal

TG/HDL-C ratio above 1.74 is too high

TG/HDL-C ratio above 2.62 is much too high

In this article, TG/HDL-C ratio is provided as in the US (mg/dl).


Results

Clinical, Biochemical, and Histologic Profile

A total of 9 subjects in each group were studied (Table 1). The 3 groups were similar with respect to gender, ethnicity, age, body mass index, fasting blood sugar, glycosylated hemoglobin, and hepatic synthetic functions. Subjects with NASH had higher aspartate aminotransferase and alanine aminotransferase levels that approached but did not reach significance. Although the total cholesterol was higher in subjects with NAFL and NASH than controls (P < 0.05 for both), there were no significant differences in high-density lipid cholesterol or low-density lipid cholesterol. Subjects with NAFL had isolated steatosis (mean grade ± SEM: 2.1 ± 0.6) or steatosis with mild inflammation (mean NAFLD activity score ± SEM: 3.2 ± 0.3), whereas those with NASH had a mean steatosis score ± SEM of 1.8 ± 0.4 and an NAFLD activity score ± SEM of 5.1 ± 0.4 (P < 0.03 versus NAFL).

Parameter Control (n = 9) NAFL (n = 9) NASH (n = 9)
Gender F/M 7/2 6/3 6/3
Caucasian/African American 7/2 8/1 9/0
Mean age (years) 46.6 ± 3.8 48.5 ± 4.8 47 ± 3.2
BMI (kg/m 2 ) 34.5 ± 4.3 37.5 ± 5.2 34 ± 1.8
Fasting glucose (mg/dl) 87.5 ± 4.9 119.2 ± 39.6 123.3 ± 29.3
Glycosylated hemoglobin (%) 5.2 ± 0.4 5.8 ± 0.1 5.8 ± 0.1
AST (normal range: 0-65 U/l) 42 ± 16.6 36.8 ± 5 77.4 ± 14.2
ALT (normal range: 0-65 U/l) 55.7 ± 27.4 49 ± 10.2 119.4 ± 25.8
Alkaline phosphatase (U/l) 82.4 ± 9.1 113.4 ± 22.8 111 ± 13.9
Total bilirubin (mg/dl) 0.6 ± 0.1 0.5 ± 0.1 0.5 ± 0.1
Albumin (g/dl) 4.1 ± 0.1 4 ± 0.3 4 ± 0.1
Platelets (1000/mm 3 ) 255.5 ± 27.9 281.3 ± 40.5 233 ± 23.3
Total cholesterol (mg/dl) 166.5 ± 10.5 230.6 ± 13.8* * P < 0.05 versus the control (analysis of variance).
234 ± 11.2* * P < 0.05 versus the control (analysis of variance).
HDL cholesterol (mg/dl) 57.5 ± 8.5 40.2 ± 6.3 50.5 ± 2.1
LDL cholesterol (mg/dl) 87 ± 4 127 ± 22.6 99 ± 3.8
Triglycerides (mg/dl) 220 ± 22 397.8 ± 135.2 274 ± 19.4
Steatosis grade 0 2.1 ± 0.6 1.8 ± 0.4
NAFLD activity score 0 3.2 ± 0.3 5.1 ± 0.4† † P < 0.05 versus NAFL (t test).
Fibrosis stage 0 1.3 ± 0.3 1.8 ± 0.4
  • The data are expressed as the mean ± SEM. ALT indicates alanine aminotransferase AST, aspartate aminotransferase BMI, body mass index HDL, high-density lipid LDL, low-density lipid NAFL, nonalcoholic fatty liver NAFLD, nonalcoholic fatty liver disease NASH, nonalcoholic steatohepatitis.
  • * P < 0.05 versus the control (analysis of variance).
  • P < 0.05 versus NAFL (t test).

Total Hepatic Lipid Content (Table 2 )

The total hepatic lipid content was markedly increased in NAFL and NASH (P < 0.001) this was driven mainly by increased TAG content (P < 0.001). Similarly, DAG was also increased significantly (P < 0.03 for both NAFL and NASH). Compared to that of normal controls, the TAG/DAG (product/precursor) ratio was significantly increased in both NAFL and NASH (7 versus 26 versus 31, P < 0.001 for both). The n-6 fatty acid content in the total lipids increased from the controls to NAFL and NASH (mean ± SEM: 4131 ± 210 versus 6424 ± 605 versus 8449± 1012 nmol/gm, P < 0.03 for NASH versus the controls) and was associated with a significantly higher total n-6:n-3 ratio in NAFL and NASH (P < 0.05 for both versus the controls). However, the FFA content did not increase in either NAFL or NASH. Also, there was a stepwise increment in FC from normal to NAFL to NASH (P < 0.05 for the control versus NASH). This was, however, not accompanied by an increase in CEs in either NAFL or NASH.

Parameter Control (n = 9) NAFL (n = 9) NASH (n = 9)
Total lipids 15,978 ± 1,157 49,991 ± 10,718* * P < 0.05 versus the control (analysis of variance).
43,658 ± 6,162* * P < 0.05 versus the control (analysis of variance).
Total SFA 7,189 ± 586 22,889 ± 5,328* * P < 0.05 versus the control (analysis of variance).
18,756 ± 3,003* * P < 0.05 versus the control (analysis of variance).
Total MUFA 3,527 ± 258 19,616 ± 4,937 14,819 ± 2,204* * P < 0.05 versus the control (analysis of variance).
Total PUFA 5,104 ± 347 7,328 ± 606* * P < 0.05 versus the control (analysis of variance).
9,822.62 ± 1,094* * P < 0.05 versus the control (analysis of variance).
Total n-3 949 ± 153 862 ± 49 1,336 ± 180
Total n-6 4,131 ± 210 6,424 ± 605 8,449 ± 1,012* * P < 0.05 versus the control (analysis of variance).
n-6/n-3 ratio 4.8 ± 0.4 7.6 ± 0.8* * P < 0.05 versus the control (analysis of variance).
6.9 ± 0.9* * P < 0.05 versus the control (analysis of variance).
FFA 5,533 ± 1,210 5,929 ± 1,487 6,115 ± 584
DAG 1,922 ± 430 4,946 ± 1,232* * P < 0.05 versus the control (analysis of variance).
3,304 ± 473* * P < 0.05 versus the control (analysis of variance).
TAG 13,609 ± 2,300 128,585 ± 34,201* * P < 0.05 versus the control (analysis of variance).
104,035 ± 20,451* * P < 0.05 versus the control (analysis of variance).
TAG/DAG ratio 7 26* * P < 0.05 versus the control (analysis of variance).
31* * P < 0.05 versus the control (analysis of variance).
CE 7,589 ± 1,548 8,522 ± 1,846 7,774 ± 808
FC 7,538 ± 718 10,382 ± 1,079 12,862 ± 1,707* * P < 0.05 versus the control (analysis of variance).
CL 4,447 ± 620 5,023 ± 585 6,063 ± 902
LYPC 1,936 ± 308 1,800 ± 505 2,239 ± 475* * P < 0.05 versus the control (analysis of variance).
PC 20,321 ± 1,098 15,322 ± 1,247* * P < 0.05 versus the control (analysis of variance).
16,874 ± 1,319
PE 14,599 ± 813 11,456 ± 494* * P < 0.05 versus the control (analysis of variance).
14,828 ± 1,359
PS 4,114 ± 577 5,295 ± 1,774 4,199 ± 706
SM 4,194 ± 587 3,773 ± 590 5,684 ± 685
  • All values are expressed as the mean (nmol/g of tissue) ± SEM. CE indicates cholesterol ester CL, cardiolipin DAG, diacylglycerol FC, free cholesterol FFA, free fatty acid LYPC, lysophosphatidylcholine MUFA, monounsaturated fatty acid NAFL, nonalcoholic fatty liver NASH, nonalcoholic steatohepatitis PC, phosphatidylcholine PE, phosphatidylethanolamine PUFA, polyunsaturated fatty acid PS, phosphatidylserine SFA, saturated fatty acid SM, sphingomyelin TAG, triacylglycerol.
  • * P < 0.05 versus the control (analysis of variance).

The total hepatic PL content was not significantly different across the 3 groups (mean ± SEM: controls = 49,889 ± 3220 nmol/g of tissue, NAFL = 42,671 ± 3714 nmol/g of tissue, and NASH = 49,614 ± 2599 nmol/g of tissue, P = not significant). However, despite a significant increase in the total lipids and FC, the total PC content was decreased (P < 0.03 for NAFL versus the control). This was accompanied by an increase in lysophosphatidylcholine (LYPC i.e., lysolecithin) in subjects with NASH (P < 0.05). There was also a trend for decreased PE that was significant only for NAFL (P < 0.05) in contrast, phosphatidylserine (PS) levels remained unchanged. Although the hepatic content of sphingomyelin (SM) and CLs increased, they did not meet levels of significance.

Fatty Acid Composition of Hepatic FFAs (FFA Table 3 )

There was a trend for a progressive decrease from controls to NAFL and then NASH for n-6 and n-3 polyunsaturated fatty acids (PUFAs) within hepatic FFA. The content of linoleic acid (18:2n-6), the starting point for processing n-6 essential fatty acids (EFAs), 18 was unaltered in both NAFL and NASH. However, there was a trend toward progressive depletion of γ-linolenic acid (18:3n-6), which is immediately downstream of linoleic acid, 18 , 19 from the control to NAFL to NASH (P < 0.01 versus NASH). This was also seen with arachidonic acid (20:4-n6), which is further downstream of γ-linolenic acid, with a significant decrease in NASH (P < 0.05 versus the controls). The product/precursor ratio for the n-6 pathway (20:4n-6:18:2n-6) trended downward, reaching significance for NASH (P < 0.03 versus the controls). The levels of mead acid (20:3n-9), which are typically increased in EFA deficiency, 20 remained unaltered in NAFL and NASH.

Fatty Acid Control (n = 9) NAFL (n = 9) NASH (n = 9)
14:0 6.6 ± 1.5 5.4 ± 0.9 5.9 ± 0.7
15:0 1.6 ± 0.3 1.2 ± 0.2 1.6 ± 0.1
16:0 30 ± 0.7 31.5 ± 1.6 32.3 ± 1.2
18:0 19.6 ± 0.9 16.8 ± 2.6 19 ± 1.8
16:1n-7 1.6 ± 0.2 2.4 ± 0.3 2 ± 0.3
18:1n-7 1.8 ± 0.3 1.8 ± 0.3 1.8 ± 0.3
18:1n-9 15.8 ± 0.8 19.2 ± 1.9 17.5 ± 1.4
24:1n-9 1.2 ± 0.3 1 ± 0.3 1.1 ± 0.1
18:2n-6 7.5 ± 0.8 8.2 ± 1.4 8.3 ± 0.8
18:3n-6 1.1 ± 0.4 0.5 ± 0.2 0.2 ± 0.1* * P < 0.05 versus the control.
20:4n-6 4.9 ± 0.8 4 ± 1.1 3.1 ± 0.4* * P < 0.05 versus the control.
20:5n-3 0.4 ± 0.1 0.3 ± 0.1 0.2 ± 0.1
22:6n-3 1.3 ± 0.3 1.1 ± 0.3 0.8 ± 0.2
SFA 59.3 ± 1.9 56.4 ± 2.4 60.4 ± 1.9
MUFA 22.2 ± 1.1 26.2 ± 1.7 24.2 ± 1.8
PUFA 18.4 ± 1.8 17.4 ± 1.7 15.3 ± 1.1
LCPUFA 6.6 ± 1.1 5.4 ± 1.4 4.1 ± 0.6* * P < 0.05 versus the control.
n-3 PUFA 3.2 ± 0.5 2.9 ± 0.3 2.2 ± 0.4
n-6 PUFA 15.1 ± 1.5 14.3 ± 1.5 12.9 ± 0.8
n-6/n-3 ratio 5.3 ± 0.8 5.1 ± 0.4 6.7 ± 0.8* * P < 0.05 versus the control.
† † P < 0.05 versus NAFL (analysis of variance).
20:4n-6/18:2n-6 ratio 0.7 ± 0.1 0.6 ± 0.2 0.4 ± 0.1* * P < 0.05 versus the control.
20:5n-3/18:3n-3 ratio 0.6 ± 0.2 0.6 ± 0.2 0.3 ± 0.1
  • All values are the mean (mol %) ± SEM. FFA indicates free fatty acid LCPUFA, long-chain polyunsaturated fatty acid (sum of 20:4n-6, 20:5n-3, and 22:6n-3) MUFA, monounsaturated fatty acid NAFL, nonalcoholic fatty liver NASH, nonalcoholic steatohepatitis PUFA, polyunsaturated fatty acid SFA, saturated fatty acid.
  • * P < 0.05 versus the control.
  • P < 0.05 versus NAFL (analysis of variance).

The α-linolenic acid (18:3n-3), the starting point for processing n-3 EFA, 18 was unaltered. There was a trend for a progressive decrease from the controls to NAFL to NASH for eicosapentanoic acid (20:5n-3) and docosahexanoic acid (22:6n-3), the downstream products of α-linolenic acid (18:3n-3), which approached but did not reach significance in NASH. The product/precursor ratio for the n-3 pathway (20:5n-3:18:3n-3) also trended downward, approaching significance for NASH.

Fatty Acid Composition of Hepatic DAG and TAG

Both DAG and TAG were significantly increased in subjects with NAFL and NASH (Fig. 1A). There was a trend for increased saturated fatty acids (SFAs) in DAG and TAG this was driven by increased palmitate (16:0) but offset by decreased stearic acid (18:0). There was also a trend for increased monounsaturated fatty acids (MUFAs Fig. 1B), specifically oleic acid (18:1-n9), in DAG and TAG for both NAFL and NASH.

Hepatic lipid composition in subjects with metabolic syndrome and normal liver histology (controls, n = 9), nonalcoholic fatty liver (NAFL, n = 9), and nonalcoholic steatohepatitis (NASH, n = 9). (A) Distribution of lipid classes within hepatic lipids. The x axis represents free fatty acid (FFA), diacylglycerol (DAG), and triacylglycerol (TAG) lipid classes. The y axes represent lipid contents in the 3 study groups (nmol/g of tissue), with FFA and DAG displayed on the left y axis and TAG on the right y axis. NAFL and NASH had significantly increased hepatic DAG (P < 0.05) and TAG (P < 0.002). The FFA content did not change in either group. (B) There was a trend of an increase in the mole percent of saturated fatty acids (SFAs) and monounsaturated fatty acids (MUFAs) within TAG in both NAFL and NASH. The polyunsaturated fatty acid (PUFA) content of TAG, however, was significantly depleted in NAFL and NASH (P < 0.01). (C) Both n-3 (left y axis) and n-6 (right y axis) PUFAs were significantly decreased in NAFL and NASH (P < 0.02). (D) The n-6:n-3 ratio was significantly higher in NAFL and NASH (P < 0.01 for both). (E,F) Relative changes from the control (shown as 100%) are plotted. The arachidonic acid (20:4n-6), eicosapentanoic acid (20:5n-3), and docosahexanoic acid (22:6n-3) were significantly depleted in both NAFL and NASH. (B-F) The data shown are mole percent data (percentage of each fatty acid of the total fatty acids within each lipid class) within the TAG class. All values are expressed as the mean ± SEM. *P < 0.05 versus the controls. The hollow bars represent the controls, the solid bars represent NAFL, and the slashed bars represent NASH.

In contrast, there was a significant decrease in PUFA associated with DAG and TAG in NAFL and NASH (Fig. 1B). The molar percentages of n-3 and n-6 fatty acids in TAG were decreased in both NAFL and NASH (Fig. 1C) however, the decrease in n-6 was less than that in the n-3 fatty acids, resulting in a net significant increase in the n-6:n-3 fatty acid proportions in TAG (Fig. 1D). There was a significant depletion of arachidonic acid (20:4n-6), a key n-6 fatty acid (Fig. 1E). Also, eicosapentanoic acid (20:5n-3) and docosahexanoic acid (22:6n-3), the 2 downstream products of α-linolenic acid (18:3n-3) in the n-3 pathway, were significantly depleted (Fig. 1F). These changes were qualitatively similar to the changes seen in the FFA and DAG pools.

Hepatic CE Fatty Acid Composition

The hepatic FC content increased progressively from controls with normal histology to NAFL to NASH (P < 0.05 for NASH versus the control Fig. 2A). The total CE content was not, however, significantly changed in either NAFL or NASH. There was a de-enrichment of SFA and a mild, insignificant increase in MUFA, specifically oleic acid. There was a significant enrichment of the CEs with PUFA (Fig. 2B,C). Both n-6 and n-3 PUFAs increased in NAFL and NASH, but the findings were significant only for n-6 fatty acids (Fig. 2C). The overall n-6:n-3 ratio did not change significantly (Fig. 2D). Although linoleic acid (18:2n-6) was particularly enriched within the CEs in both NAFL and NASH, the arachidonic acid levels were not altered significantly (Fig. 2E). Although there was a trend of an increase in docosahexanoic acid (22:6n-3) in NASH, this did not reach significance (Fig. 2F).

Hepatic cholesterol composition in controls (n = 9), nonalcoholic fatty liver (NAFL n = 9), and nonalcoholic steatohepatitis (NASH n = 9). Parts B-F reflect changes within the cholesterol ester (CE) class and are from mole percent data (percentage of each fatty acid of the total fatty acids within each lipid class). (A) Distribution of CE and free cholesterol (FC) within hepatic lipids. There was a stepwise increase in the hepatic FC content from the controls to NAFL to NASH (P < 0.05, NASH versus the control). The CE content was comparable in the 3 groups. (B) The hepatic CE was significantly depleted in saturated fatty acids (SFAs P < 0.02) and enriched with polyunsaturated fatty acids (PUFAs P < 0.01) in both NAFL and NASH. (C) Although both n-3 (left y axis) and n-6 (right y axis) PUFAs were increased in NAFL and NASH, it was significant only for n-6 (P < 0.02). (D) The n-6:n-3 ratio was not significantly different among the 3 groups. (E,F) The relative changes from the control (shown as 100%) are plotted. The linoleic acid (18:2n-6) content was significantly higher (P < 0.01), with a trend of a decrease in arachidonic acid (20:4n-6) and a trend of an increase in docosahexanoic acid (22:6n-3) in both NAFL and NASH. All values are expressed as the mean ± SEM. *P < 0.05 versus the controls. The hollow bars represent the controls, the solid bars represent NAFL, and the slashed bars represent NASH.

Fatty Acid Composition of Hepatic PLs

The total PC content in the liver was decreased in both NAFL and NASH, but no significant changes were seen in PS or CL (Fig. 3A,B). There was an accompanying increase in LYPC (P < 0.05 for NASH versus the control Fig. 3A). The mole percentages of SFA, MUFA, and PUFA were unchanged in PC (Fig. 3C). There were also no significant changes in total n-6 or n-3 fatty acids in PC (Fig. 3D). Although the n-6 EFA linoleic acid (18:2n-6) content was unaltered, there was a progressive stepwise decrease in arachidonic acid (20:4n-6) that was significant for NASH (P < 0.05 versus the controls Fig. 3E). The resultant decrease in the product/precursor ratio for the n-6 pathway (20:4n-6:18:2n-6) reached significance for NASH (P < 0.01 versus the controls).

Hepatic phospholipid composition in controls (n = 9), nonalcoholic fatty liver (NAFL n = 9), and nonalcoholic steatohepatitis (NASH n = 9). Parts C-F show changes within phosphatidylcholine (PC) based on mole percent data (percentage of each fatty acid of the total fatty acids within each lipid class). (A) The PC content in liver was decreased in both NAFL and NASH. This was associated with a significant increase in lysophosphatidylcholine (LYPC P < 0.05) and a trend of an increase in hepatic sphingomyelin (SM right y axis) only in NASH. (B) No significant changes were noted in phosphatidylserine (PS) and cardiolipins (CLs), with decreased phosphatidylethanolamine (PE) in NAFL. (C) No alterations were found in the saturated fatty acid (SFA), monounsaturated fatty acid (MUFA), and polyunsaturated fatty acid (PUFA) contents of hepatic PC. (D) The n-3 (left y axis) and n-6 (right y axis) PUFAs were unchanged in NAFL and NASH. (E,F) The relative changes from controls (shown as 100%) are plotted. Arachidonic acid (20:4n-6) in hepatic PC was significantly depleted in NASH (P < 0.01). There was also a trend of a decrease in eicosapentanoic acid (20:5n-3) and docosahexanoic acid (22:6n-3) in both NAFL and NASH. All values are expressed as the mean ± SEM. *P < 0.05 versus the controls. The hollow bars represent the controls, the solid bars represent NAFL, and the slashed bars represent NASH.

The α-linolenic acid (18:3n-3) content of PC was unchanged in NAFL and NASH (Fig. 3F). There was, however, a trend for decreased downstream n-3 fatty acid products, specifically eicosapentanoic acid (20:5n-3) and docosahexanoic acid (22:6n-3). This trend was in keeping with the profile seen in TAG and DAG and in the FFA pool in the liver. Also, in keeping with the profile seen in these lipid classes, there was a progressive, but nonsignificant, trend of an increase in the total n-6:n-3 ratio in PC from 6.7 in normal controls to 7.6 in NAFL and 8.5 in NASH.

Changes in Essential n-6 and n-3 Fatty Acids

The n-6 fatty acid mole percentage was depleted in FFA, DAG, and TAG but reached significance for only TAG in both NAFL and NASH (Table 4). Arachidonic acid (20:4n-6) was relatively depleted from most lipid classes. Specifically, there were 31% and 36% decreases in the arachidonic acid (20:4n-6) content of PC (P < 0.03) and DAG (P = not significant) in NASH, respectively, the principal sources for its release for inflammatory prostaglandin synthesis. 21 This decrease was noted despite no significant changes in its precursor linoleic acid (18:2n-6) in any of these classes.

Lipid Class n-3 PUFA n-6 PUFA n-6:n-3 Ratio
Control NAFL NASH Control NAFL NASH Control NAFL NASH
FFA 3.1 ± 0.5 2.8 ± 0.3 2.2 ± 0.4 15.1 ± 1.5 14.3 ± 1.5 12.9 ± 0.8 5.2 ± 0.8 5.1 ± 0.4 6.7 ± 0.8* * P < 0.05 versus the control.
† † P < 0.05 versus NAFL (analysis of variance).
DAG 4.0 ± 1.3 2.1 ± 0.4 2.1 ± 0.4 17.5 ± 1.9 11.8 ± 1.2* * P < 0.05 versus the control.
16.1 ± 0.6 6 ± 0.9 6.4 ± 1.2 10.3 ± 2.5
TAG 1.9 ± 0.3 0.6 ± 0.1* * P < 0.05 versus the control.
1.2 ± 0.1* * P < 0.05 versus the control.
18.8 ± 0.7 10.8 ± 1.7* * P < 0.05 versus the control.
15.5 ± 1.1* * P < 0.05 versus the control.
10.2 ± 0.9 16.1 ± 1.4* * P < 0.05 versus the control.
13.2 ± 0.8* * P < 0.05 versus the control.
CE 1.5 ± 0.4 2.2 ± 0.2 2.7 ± 0.5 17.4 ± 1.9 28.2 ± 4.1* * P < 0.05 versus the control.
26.3 ± 2.8* * P < 0.05 versus the control.
16 ± 3.1 13.5 ± 2.4 12.8 ± 2.6
CL 6.4 ± 0.7 5.6 ± 0.5 8 ± 2.1 32.8 ± 3.5 29.3 ± 3.4 29.9 ± 1.8 5.4 ± 0.7 5.4 ± 0.5 5.6 ± 1.3
LYPC 2.5 ± 0.5 2.8 ± 0.3 2.8 ± 0.3 10.8 ± 1.2 12.9 ± 0.9 12.4 ± 0.9 5.5 ± 1 4.7 ± 0.4 4.85 ± 0.6
PC 5.6 ± 0.8 4.5 ± 0.4 4.8 ± 0.6 34.5 ± 0.8 32.5 ± 1.7 33.9 ± 0.9 6.7 ± 0.7 7.6 ± 0.8 8.5 ± 1.6
PE 12.9 ± 1.4 11 ± 1.1 12 ± 1.2 32.8 ± 1.3 33.3 ± 1.5 31.3 ± 1.3 2.8 ± 0.4 3.2 ± 0.3 2.9 ± 0.4
PS 8.6 ± 1.9 8.7 ± 1.5 8.6 ± 1.1 18.3 ± 2.3 20.1 ± 2.8 16.6 ± 1.5 3.5 ± 1.2 2.4 ± 0.2 2 ± 0.1
SM 3.2 ± 0.3 2.3 ± 0.6 2.8 ± 0.6 11.1 ± 0.8 10.6 ± 1.4 10.2 ± 1.3 3.5 ± 0.1 5.8 ± 1.3 4.8 ± 0.9
Arachidonic Acid (20:4n-6) Eicosapentanoic Acid (20:5n-3) Docosahexanoic Acid (22:6n-3)
FFA 4.8 ± 0.8 4 ± 1.1 3.1 ± 0.3* * P < 0.05 versus the control.
0.41 ± 0.1 0.3 ± 0.1 0.2 ± 0.1 1.3 ± 0.3 1.1 ± 0.3 0.8 ± 0.2
DAG 4.8 ± 1 2.2 ± 0.5 3.1 ± 0.5 0.22 ± 0.1 0.2 ± 0.1 0.2 ± 0.1 1.1 ± 0.3 0.6 ± 0.2 0.7 ± 0.2
TAG 1.3 ± 0.2 0.3 ± 0.1* * P < 0.05 versus the control.
0.3 ± 0.1* * P < 0.05 versus the control.
0.14 ± 0.04 0.02 ± 0.01* * P < 0.05 versus the control.
0.05 ± 0.02* * P < 0.05 versus the control.
0.5 ± 0.1 0.06 ± 0.01* * P < 0.05 versus the control.
0.2 ± 0.05* * P < 0.05 versus the control.
CE 4.2 ± 0.4 3.9 ± 0.7 3.2 ± 0.4 0.24 ± 0.1 0.3 ± 0.1 0.3 ± 0.1 0.4 ± 0.1 0.6 ± 0.2 1.0 ± 0.4
CL 7.6 ± 0.5 7.4 ± 1.0 6.3 ± 0.4 0.17 ± 0.03 0.3 ± 0.1 0.3 ± 0.1 5 ± 0.7 4.1 ± 0.4 5.5 ± 1.9
LYPC 4.6 ± 0.9 5.7 ± 0.8 4.5 ± 0.7 0.21 ± 0.1 0.4 ± 0.1 0.2 ± 0.1 1.3 ± 0.3 1.4 ± 0.3 1.6 ± 0.3
PC 12.3 ± 1.2 9.5 ± 1.1 8.5 ± 0.6* * P < 0.05 versus the control.
1.0 ± 0.3 0.9 ± 0.2 0.8 ± 0.1 3.5 ± 0.5 2.5 ± 0.2 2.9 ± 0.5
PE 22.4 ± 1.1 21.0 ± 1.4 18.8 ± 0.8 0.6 ± 0.2 0.8 ± 0.2 0.9 ± 0.3 10.8 ± 1.4 8.5 ± 0.9 9.5 ± 1.3
PS 10.3 ± 2.2 11.2 ± 2.4 8.8 ± 1 0.4 ± 0.1 0.6 ± 0.1 0.4 ± 0.1 6.3 ± 1.5 6.2 ± 1.5 6.6 ± 1.0
SM 3.6 ± 0.4 3.2 ± 0.7 2.7 ± 0.6 0.2 ± 0.1 0.1 ± 0.1 0.2 ± 0.1 1.5 ± 0.5 1.4 ± 0.4 1.5 ± 0.5
  • All values are the mean (mol%) ± SEM. CE indicates cholesterol ester CL, cardiolipin DAG, diacylglycerol FFA, free fatty acid LYPC, lysophosphatidylcholine NAFL, nonalcoholic fatty liver NASH, nonalcoholic steatohepatitis PC, phosphatidylcholine PE, phosphatidylethanolamine PUFA, polyunsaturated fatty acid PS, phosphatidylserine SM, sphingomyelin TAG, triacylglycerol.
  • * P < 0.05 versus the control.
  • P < 0.05 versus NAFL (analysis of variance).

The mole percent of n-3 fatty acids was decreased in FFA, DAG, TAG, and PC in NASH but was significant only for TAG (Table 4). Eicosapentanoic acid (20:5n-3) and docosahexanoic acid (22:6n-3) were significantly depleted in TAG, and a trend of depletion was seen in FFA, DAG, and, to a lesser degree, PC. These changes occurred despite the unaltered content of the precursor n-3 EFA α-linolenic acid (18:3n-3). There was a trend of an increase in the n-6:n-3 ratio across several lipid classes, but it reached significance only for FFA (28% NASH versus both groups, P < 0.05) and TAG (57% NAFL and 29% NASH, versus the control, P < 0.05). Long-chain PUFAs (sum of 20:4n-6, 20:5n-3, and 22:6n-3) were significantly depleted in NASH (P < 0.05 versus the control) within FFA.


3 RESULTS

3.1 The proteome profiling differences between NAFLD-driven HCC tumor tissues and adjacent non-tumor tissues

In total, 1965 proteins were identified and quantified (Table S5). To avoid potential mis-identification bias, we focused on 238 proteins that were identified in all samples. A volcano plot was generated to show the differential expression distribution (Figure 1A). Among the 238 proteins, 100 proteins presented significant differential expression between HCC tissues and adjacent non-tumor tissues (P < .05) (Table S6). Here, 65 proteins were upregulated and 35 proteins were downregulated in the HCC. Pathway analysis indicated that the top enriched pathways included Ketogenesis, Mevalonate Pathway I and Isoleucine Degradation I (Figure 1B and Table S7). The significant altered proteins were analyzed by using the PANTHER classification system (http://pantherdb.org/). 22 The majority of proteins belonged to cell components (58.68%), possessed catalytic activity (45.33%), and were involved in metabolic processes (31.77%) (Figure S1). Pathway analysis was consistent with findings in a previous report of an increased glycolysis process and a decreased fatty acid beta-oxidation process. Furthermore, our proteomics results were comparable with those of a previous study, indicating that a potential mechanism difference existed between NAFLD-associated HCC and other types of HCC. 23-25 The results of early-stage hepatocellular carcinoma proteomics showed that hepatocellular carcinoma was related to hepatitis B virus infection (HBV-HCC) and was associated with disrupted cholesterol homeostasis, accompanied by high expression of sterol O-acyltransferase 1 (SOAT1). 23 Nonetheless, our study suggested that the differences in protein expression between NAFLD-driven HCC tissues and adjacent non-tumor tissues were mainly focused on ketogenesis. This difference suggested that different diagnoses and therapeutic strategies should be applied to different types of HCC.

3.2 VDAC1 is dysregulated in NAFLD-driven HCC and associated with NAFLD

To prioritize the genes associated with NAFLD, a transcriptome-wide association analysis was performed in BXD mice strains. The correlation between body fat mass percentage and the transcripts of 100 candidate proteins was calculated. Among all candidates, VDAC1 presented a significant correlation with body fat mass percentage (R = 0.469, P = .008) (Figure 1C and Table S1). To further confirm the association between VDAC1 and NAFLD, the correlation between VDAC1 and a series of fatty liver disease phenotypes was evaluated at both the transcript and peptide levels. The phenotypes included body fat mass, body weight, liver mass, white adipose mass, cholesterol in plasma, basal glucose in plasma under fasting and area under curve of glucose in an oral glucose tolerance test (Glucose AUC in OGTT) (Figure 1E and Table S2). The results suggested that both the gene and protein expression levels of VDAC1 were correlated with NAFLD traits.

Dysregulation of VDAC1 in HCC was further validated. Our proteomic results suggested a 3.3-fold increase in VDAC1 in HCC tissues. This result was validated using a western blot (Figures 1D and S2), and further confirmed in an external clinical cohort. In The Cancer Genome Atlas (TCGA) LIHC cohort, Vdac1 expression levels were upregulated in HCC tissues compared with non-tumor tissues (Normal N = 50, Tumor N = 371, P < .0001) (Figure 2A) and increased with increasing tumor grade (Figure 2B). Moreover, high-level VDAC1 expression was correlated with poor clinical outcome (High, n = 91 Low, n = 91 P = .013) (Figure 2C). Collectively, these results indicated that VDAC1 was associated with NAFLD and dysregulated NAFLD-driven HCC.

3.3 System genetics analysis of Vdac1 in BXD mice strains

System genetics analysis was then performed to reveal the expression regulation of VDAC1. Gene expression levels of VDAC1 in liver tissue 39 BXD mouse strains and their corresponding parental strains were examined. There was only 1 probe set (10 375 941) that targeted the first exon of the Vdac1 gene in the Affymetrix Mouse Gene 1.0 ST array. Vdac1 expression varied widely across BXD strains with a fold change of 1.41 (Figure 3A). The average expression of Vdac1 across all BXD strains was 12.12 ± 0.02 (log2 scale, mean ± standard error of the mean (SEM)). The strain with the highest level of expression was BXD45, while the strain with the lowest expression was BXD61. To identify sequence variants that affected the expression of Vdac1 in mouse liver, we performed simple interval mapping for Vdac1 across the mouse genome. One significant expression QTL (eQTL) was identified on chromosome 11 at 52.01 Mb with an LRS of 18.3, and which was close to the physical location of Vdac1 on chromosome 11 at 52.36 Mb (Figure 3B and Table S8). In addition, we also identified 4 suggestive eQTLs located on other chromosomes (Figure 3B and Table 2). These results indicated that the Vdac1 mRNA level was both cis- and trans-regulated, simultaneously. For the other 4 trans-eQTLs, we retrieved a total of 361 protein coding genes from their 1.5 LOD confidence interval (Table 2). Among them, 18 genes are significantly correlated with the Vdac1 expression level (Table 2). From the liver proteome data, there were 20 peptide-targeted VDAC1 proteins. To evaluate the correlation between Vdac1 transcripts and peptides, a correlation analysis was performed. Among the 20 peptides, the top 5 peptides most highly correlated with the VDAC1 transcript were listed, Peptide 3 (R = 0.458, P = .005), Peptide 10 (R = 0.472, P = .004), Peptide 14 (R = 0.493, P = .002), Peptide 15 (R = 0.437, P = .008), and Peptide 19 (R = 0.496, P = .002) (Figure S3 and Table S4). Therefore, Peptides 3, 10, 14, 15, 19 were chosen to represent the VDAC1 protein for further analysis.

Chr Start End LRS Marker # of gene Genes correlated with Vdac1
2 98 107 16.263 rs33188980 87 Tcp11l1 Apip Cd59a Cd59b Lmo2 Ldlrad3 Eif3m Qser1
5 132 136 14.215 rs36653585 66 Lat2 Limk1 Mdh2
14 73 100 12.941 151 Klf5 Rcbtb2 Tpt1 Mzt1 Sucla2 Esd
X 64 72 16.188 rs33876026 57 Prrg3

3.4 Gene co-expression network and function enrichment analysis indicated VDAC1 is associated with mitochondria dysfunction

To explore the potential mechanism of VDAC1 involved in NAFLD-driven HCC, the Vdac1 gene co-expression network and functional enrichment analysis were implemented. With the aim to identify Vdac1 genes involved in gene modules, the filtered probes were submitted to the WGCNA 18 package for constructing gene co-expression networks. Soft threshold power β (8) was chosen based on scale-free topology criteria (Figure 4A,B). Using this soft-thresholding power, we identified 35 distinct co-expression modules that were assigned different colors Vdac1 was located in the green module containing 589 genes (Figure 4C and Table S9). To verify the involvement of the green module in the biological processes, we performed GO and KEGG enrichment analysis using the WebGestalt online toolkit. We found that this module was significantly enriched in biological processes such as the oxidation-reduction process, cell death, and the apoptotic signaling pathway (Table 3). This green module was also enriched in several KEGG pathways including oxidative phosphorylation, NAFLD, and the tricarboxylic acid cycle (TCA cycle) (Table 3). The analysis revealed that Vdac1 and its co-expressed genes present in the same module were involved in biological functions such as energy metabolism and liver disease (NAFLD). Furthermore, pathway analysis using IPA showed that the mitochondrial dysfunction pathway was the top enriched pathway (Figure 5A and Table S10) and that 20 mitochondrial inner membrane proteins were involved in this pathway, which showed correlation with Vdac1 (Figure 5B and Table S11).

Term Description No. of Genes P-value FDR
GO: Biological process
GO:0055114 Oxidation-reduction process 54 4.89E-10 4.08E-06
GO:0045333 Cellular respiration 18 4.17E-09 1.74E-05
GO:0006091 Generation of precursor metabolites and energy 24 6.68E-08 1.85E-04
GO:0015980 Energy derivation by oxidation of organic compounds 20 2.06E-07 2.86E-04
GO:0007005 Mitochondrion organization 29 4.36E-06 2.45E-03
GO:0008219 Cell death 67 4.42E-06 2.45E-03
GO:0097190 Apoptotic signaling pathway 30 5.72E-06 2.75E-03
GO:0006915 Apoptotic process 62 1.04E-05 3.61E-03
GO:0012501 Programmed cell death 62 1.63E-05 4.84E-03
GO:0006099 Tricarboxylic acid cycle 6 3.27E-05 7.79E-03
KEGG pathways
mmu00190 Oxidative phosphorylation 22 2.17E-12 6.49E-10
mmu05012 Parkinson disease 21 7.31E-11 1.09E-08
mmu05010 Alzheimer disease 21 2.91E-09 2.90E-07
mmu05016 Huntington disease 21 1.88E-08 1.40E-06
mmu01100 Metabolic pathways 61 8.03E-07 4.80E-05
mmu04932 Non-alcoholic fatty liver disease (NAFLD) 16 1.60E-06 7.98E-05
mmu01200 Carbon metabolism 13 8.85E-06 3.78E-04
mmu00020 Citrate cycle (TCA cycle) 6 1.34E-04 5.02E-03
mmu04142 Lysosome 10 1.23E-03 4.09E-02
mmu00010 Glycolysis/Gluconeogenesis 7 1.39E-03 4.16E-02

3.5 VDAC1 is correlated with cardiolipin (CL) profiling shift

To further understand how VDAC1 was associated with mitochondrial dysfunction, we investigated the association between VDAC1 and mitochondrial membrane lipids. CL was the signature phospholipid of the IMM, which plays an important role in maintaining the structure and function of mitochondria. 26, 27 A gene-lipidome association analysis was performed between VDAC1 and 22 lipid species from the CL class. Expression levels of the VDAC1 transcript and peptides had a negative correlation with the mature CLs, CL (LLLL), and its precursor MLCL (LLL) (Figure 5C and Table S3). Conversely, VDAC1 had a positive correlation with the nascent CLs panel (Figure 5C and Table S3), thus we hypothesized that VDAC1 was associated with CL remodeling impairment. Therefore, we calculated the nascent CLs/mature CL species CL(LLLL) ratio for each nascent CL species that is the hallmark of CL remodeling. VDAC1 was positively correlated with 17 out 18 nascent/mature ratios (Table S12). Moreover, the significant correlation between VDAC1 and cardiolipin synthetase PTPMT1 and TAZ was confirmed in the BXD mice cohort with (R = 0.452, P = .003 R = 0.334, P = .033) (Figure 5D and Table S13). VDAC1 was correlated with CLs synthesis and CL remodeling, which might damage the stability of mitochondrial inner membrane structure by changing the CL components, and eventually cause mitochondrial dysfunction.

3.6 Mitochondrial function analysis with or without VDAC1 expression in Hep3B and HepG2

To further identify whether VDAC1 overexpression attenuates mitochondrial function or not, we performed pathological and biological experiments. The results of immunohistochemical analysis suggested that the expression of VDAC1 elevated in NAFLD-driven HCC tissues compared with that in the adjacent non-tumor tissues (Figure 6E). To illustrate the effects of VDAC1 on mitochondrial function, HCC cells (HepG2 and Hep3B) were applied to knockdown or overexpress VDAC1. Meanwhile, we also applied an inhibitor to reduce VDAC1 function. VDAC1 reduction and overexpression were determined by western blot (Figure 6A). After depletion of VDAC1 in HCC cells, the mitochondrial respiratory function enhanced significantly compared with the overexpression VDAC1 group (Figure 6D). Furthermore, overexpression of VDAC1 significantly affected mitochondrial respiration, with inhibition of oxygen consumption rates (OCRs) in basal respiration (Figure 6D). The IMM with abundant respiratory enzymes played an important role in mitochondrial respiration. Additionally, the inner membrane is rich in an unusual phospholipid, cardiolipin. Subsequently, cardiolipins were quantified by fluorometric probe kit and NAO staining. Cardiolipin content was decreased by overexpression of VDAC1 in both HepG2 and Hep3B cell lines. In contrast, decreased VDAC1 caused an increase in the cardiolipin content (Figure 6B,C). These data revealed an important role of VDAC1 overexpression in the decline in mitochondrial function.

3.7 VDAC1-centralized transcript-lipids-phenotypes correlation network

To better illustrate the association of VDAC1 to NAFLD phenotypes through CL profiling, a VDAC1-centralized transcript-lipids phenotypes correlation network was generated to provide a reference for future studies (Figure 7A and Tables S2-S4). In summary, our study indicated that VADC1 was associated with NAFLD and the dysregulation of VDAC1 in NAFLD-driven HCC through mitochondrial dysfunction and by regulating mitochondrial membrane proteins and the CL acyl chain composition shift (Figure 7B).


INTRODUCTION

Dynamin-superfamily GTPases catalyze multiple membrane remodeling events in the eukaryotic cell, including those of organellar division and fusion (Praefcke and McMahon, 2004 Heymann and Hinshaw, 2009). Dynamin-related protein 1 (Drp1), a cytosolic member of this extensive superfamily, establishes the critical balance of mitochondrial division and fusion by catalyzing mitochondrial membrane fission (Chan, 2012). In response to a variety of physiological cues, Drp1 translocates to the mitochondrial outer membrane (MOM) surface to initiate the membrane remodeling events that culminate in mitochondrial division. However, the molecular mechanisms underlying these processes are just beginning to be understood.

Unlike prototypical dynamin, Drp1 lacks an apparent lipid-binding domain and requires membrane-integrated adaptor proteins such as Fis1, Mff, and MiD49/51 for its recruitment to the MOM (Bui and Shaw, 2013 Labbe et al., 2014 Richter et al., 2015). However, we and others have recently identified the mitochondrion-specific lipid cardiolipin (CL) as a specific binding partner for Drp1 (Bustillo-Zabalbeitia et al., 2014 Macdonald et al., 2014 Ugarte-Uribe et al., 2014). CL facilitates the self-assembly of Drp1 into helical polymers that tubulate membranes and also robustly stimulates Drp1 GTPase activity, two features considered functional hallmarks of these atypical GTPases (Heymann and Hinshaw, 2009). However, the nature of Drp1–CL interactions and the role of CL, if any, in the mitochondrial membrane–remodeling process remain unclear.

CL is an unusual dimeric phospholipid with unique biochemical and biophysical properties (Schlame et al., 2005 Huang and Ramamurthi, 2010). Containing four predominantly linoleoyl (C18:2) acyl chains in higher eukaryotes, native CL has a small head-group cross-sectional area and volume relative to its acyl chains, rendering it effectively cone shaped. Consequently, CL exhibits complex phase behavior in membranes, with a propensity to stabilize negative membrane curvature upon sequestration and to transition from a lamellar, bilayer phase to the nonlamellar, inverted hexagonal (HII) configuration upon further enrichment (de Kruijff and Cullis, 1980 Seddon, 1990 Tarahovsky et al., 2000 Trusova et al., 2010 Renner and Weibel, 2011). This property of CL may be relevant to lipid rearrangement events that precipitate membrane fission or fusion, especially when considered in the context of mitochondrial membrane dynamics (Chernomordik and Kozlov, 2008 Doan et al., 2013 Pan et al., 2014). Indeed, the mitochondrial inner membrane (MIM) dynamin-related GTPase OPA1 depends critically on CL for promoting mitochondrial membrane fusion (DeVay et al., 2009 Ban et al., 2010). Furthermore, defects in the acyl chain remodeling of CL and the resultant variations in CL acyl chain composition lead to the cardioskeletal myopathy Barth syndrome, underlining the importance of this atypical lipid to proper mitochondrial function (Schlame and Ren, 2006). How Drp1 and CL function cooperatively in effecting mitochondrial membrane remodeling and fission remains unknown.

In this study, we demonstrate that Drp1 stably associates with CL localized at a high spatial density in the target lipid bilayer. We further show that Drp1 preferentially reorganizes unconstrained (i.e., non–raft-associated) fluid-phase CL to generate condensed membrane platforms for subsequent membrane remodeling. Stimulated GTP hydrolysis upon helical self-assembly induces further CL rearrangement, which appears to increase the propensity of the lipid to undergo a localized lamellar-to-nonlamellar phase transition. These CL rearrangements, dependent on Drp1 B-insert–membrane interactions, create narrowly constricted membrane regions that are predisposed for fission. Thus Drp1 and CL function cooperatively in facilitating membrane remodeling and fission during mitochondrial division.


Formation and Regulation of Mitochondrial Membranes

Mitochondrial membrane phospholipids are essential for the mitochondrial architecture, the activity of respiratory proteins, and the transport of proteins into the mitochondria. The accumulation of phospholipids within mitochondria depends on a coordinate synthesis, degradation, and trafficking of phospholipids between the endoplasmic reticulum (ER) and mitochondria as well as intramitochondrial lipid trafficking. Several studies highlight the contribution of dietary fatty acids to the remodeling of phospholipids and mitochondrial membrane homeostasis. Understanding the role of phospholipids in the mitochondrial membrane and their metabolism will shed light on the molecular mechanisms involved in the regulation of mitochondrial function and in the mitochondrial-related diseases.

1. Introduction

Mitochondria are involved in a wide range of cellular processes of importance for cell survival. The inner mitochondrial membrane is the active site for the electron transport chain and ATP production. Its integrity is crucial for mitochondrial function and depends on the supply of proteins and phospholipids. As one of the major classes of lipids in the lipid bilayer of cell and organelle membranes, phospholipids are responsible for maintaining both the structural integrity of a cell and spatial separation of subcellular compartments. The major classes of phospholipids found in the mitochondrial membrane are similar to other membranes such as phosphatidylcholine (PC) and phosphatidylethanolamine (PE), and some are exclusively components of mitochondrial membrane such as cardiolipin (CL) [1].

The interaction between phospholipids and proteins is important particularly in the inner mitochondrial membrane. A significant proportion of inner membrane-associated proteins are comprised of proteins involved in the oxidative phosphorylation and their activity depends on the phospholipid composition of the membrane. Changes in the phospholipid composition can affect mitochondrial respiration [2], which has been linked to a variety of human diseases such as Barth syndrome, ischemia, and heart failure [3, 4]. The phospholipid diversity in the mitochondrial membrane is also influenced by variation in length and degree of unsaturation of fatty acyl chain present within each class of phospholipid [5]. However the role of acyl chain composition of phospholipids in mitochondrial function is still poorly understood.

The maintenance of the phospholipid composition in the mitochondrial membranes is essential for mitochondrial function, structure, and biogenesis and relies on the metabolism of phospholipids, transport into mitochondria, and supply of lipids from the diet. In this review, we focus on the phospholipid biosynthesis, trafficking and degradation, and their regulation/remodeling by dietary lipids as well as their role in inner mitochondrial membrane integrity and function.

2. Lipid Composition of Mitochondrial Membranes

Mitochondria have a structure distinct from that of other organelles since they contain two membranes: the outer mitochondrial membrane (OMM) and the inner mitochondrial membrane (IMM), which separates the intermembrane space (IMS) from the matrix. The composition of the mitochondrial membranes is similar to that of other membranes, however. The major phospholipids in the mitochondrial membranes are phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylinositol (PI), phosphatidylserine (PS), and phosphatidic acid (PA), as to plasma membrane and phosphatidylglycerol (PG) and cardiolipin (CL), are exclusively components of mitochondrial membrane (Figure 1). PC and PE are the most abundant phospholipids, comprising 40 and 30% of total mitochondrial phospholipids, respectively. PA and PS comprise 5% of the total mitochondrial phospholipids [6, 7]. Unlike plasma membrane, mitochondrial membranes contain high levels of cardiolipin (

15% of total phospholipids) and low levels of sphingolipids and cholesterol [8, 9].


(a)
(b)
(c)
(d)
(e)
(a)
(b)
(c)
(d)
(e) Structural formulas of key mitochondrial phospholipids. (a) Phosphatidylcholine—PC (b) phosphatidylethanolamine—PE (c) phosphatidic acid—PA (d) phosphatidylserine—PS (e) cardiolipin—CL.

The phospholipid CL not only has a role in maintaining membrane potential and architecture of the inner mitochondrial membrane, but also provides essential structural and functional support to several proteins involved in mitochondrial respiration. The unique structure of CL (Figure 1), which contains three glycerol backbones and four fatty acyl chains, makes it highly important for optimal function of the mitochondria [10]. The incorporation of four linoleic acid side chains in the same CL molecule renders it the primary target of the attack by free radicals, causing its peroxidation. Mitochondrial CL peroxidation appears to be an early event preceding the intrinsic apoptotic cell death [11]. Because its role in apoptosis and mitochondrial function, CL levels and CL peroxidation have been implicated in several human diseases such as atherosclerosis [12], cancer [13], Barth syndrome [14], and neurodegenerative disorders including Alzheimer’s [15] and Parkinson’s disease [16].

Another peculiarity of the IMM is that the amount of membrane-associated proteins is high. Some studies estimate the protein-lipid ratio of the inner membrane to be as high as 3 : 1 [17]. Among the IMM proteins, a significant portion of proteins comprises the oxidative phosphorylation system, which contains five multiprotein complexes. The importance of this system relies on its role in providing the cellular ATP molecules necessary for cell survival. The activity and stability of the oxidative phosphorylation proteins are affected by their interaction with phospholipids in the IMM. For example, the lack of cardiolipin in IMM was found to destabilize the complexes III and IV of the oxidative phosphorylation system [18, 19], illustrating the importance of CL for mitochondrial respiration. In addition, CL is also involved in the import and assembly of proteins. The majority of mitochondrial proteins are nuclearly encoded and require protein translocases in the mitochondrial membranes in order to be imported [20]. Several studies have shown that CL is essential for assembly and function of the translocases, allowing the integrity of the mitochondrial import machinery [21–23].

In addition to having a role in regulating protein import and activity, CL, as well as PE, is important for tubular mitochondrial morphology and membrane fusion [24–28]. Mitochondrial morphology depends on a balance between fusion and fission events. Mitochondrial fusion requires a fusogenic lipid PA which is generated by hydrolysis of CL by phospholipase D [26]. PE is another phospholipid required for the mitochondrial fusion. Disruption of the PE synthesis through PS decarboxylation pathway causes mitochondria fusion defects [25]. CL and PE accumulation within mitochondria are regulated by Ups1 and Ups2 proteins, respectively, which are involved in the phospholipid intramitochondrial trafficking (see Section 6). Ups1 and Ups2 affect the processing of OPA1 (or Mgm1 in yeast), an inner membrane dynamin-like GTPase that regulates membrane fusion [8]. It was suggested that impaired processing of Mgm1 could explain the defects in mitochondrial morphology with an altered membrane phospholipid composition [8]. A decrease in mitochondrial content of PE was found to alter mitochondrial morphology in yeast [27] and mammalian cells [25]. Joshi and colleagues have shown that CL and PE have overlapping functions in the mitochondrial fusion and that disruption of both PE and CL causes loss of mitochondrial membrane potential, and fragmentation of yeast mitochondria [27]. Fragmented mitochondria are associated with cardiomyopathy and Barth syndrome, showing the relevance of CL and PE role in the mitochondrial fusion. In this line, a yeast crd1 mutant that does not synthetize CL exhibits multiple mitochondrial and cellular defects, including loss of mitochondrial DNA, decreased respiratory function and membrane potential and reduced cell viability at elevated temperatures as a consequence of the lack of CL [29]. Similar phenotypes were found in psd1 yeast mutant, which has reduced mitochondrial PE [30]. Taken together, mitochondrial phospholipids CL and PE both regulate mitochondrial fusion/fission and biogenesis, which in turn are linked to the cellular homeostasis.

The phospholipid PA has important functions as a lipid anchor to recruit proteins involved in trafficking [31], lipid signaling [32], and fusion [33] to membrane surfaces. The regulation of mitochondria fusion and fission by PA was found to enable mitochondria to alter their morphology, to increase the efficiency of energy production, and to mark unhealthy mitochondria for autophagy [34]. In addition, PA can serve as substrate for production of the signaling lipids diacylglycerol (DAG) and lysophosphatidic acid. Mutations in the Lipin-1 gene, the PA phosphatase that generates DAG from PA, have been associated with metabolic and neurological diseases [35].

These evidences demonstrate the importance of the mitochondrial membrane phospholipids for the IMM architecture, the apoptosis, the activity of respiratory proteins, and the transport of proteins into the mitochondria. Because of their broad role in the mitochondrial function, phospholipid alterations have been associated with several diseases [36]. For example, cancer cells, which are characterized by alterations in bioenergetics and apoptosis, have altered mitochondrial phospholipid composition [37]. Cardiac pathologies result in a reduction of tetralinoleyl-CL, with consequent increase in hydrogen peroxide by the respiratory chain and apoptosis [38, 39]. Metabolic diseases, such as diabetes and nonalcoholic fatty live disease, have also been associated with decrease in mitochondrial CL and alteration of CL acyl chain remodeling [40, 41]. Another important aspect related to phospholipid composition is their change in aging. Studies have reported that aging is inversely related to the content of unsaturated phospholipids [42]. Loss of membrane fluidity resulting from lipid peroxidation, as well as decrease in mitochondrial CL content, and altered activity of the respiratory chain are also features of aging [43]. Therefore, the regulation of phospholipid homeostasis in mitochondrial membranes through their synthesis/degradation and transport into and out of the mitochondrial membrane plays a crucial role in maintaining cellular viability and health.

3. The Biosynthesis of the Major Membrane Phospholipids Occurs in the ER

Two major membrane bilayer phospholipids PC and PE are produced from choline and ethanolamine and the common lipid intermediate diacylglycerol in the process known as the de novo CDP-choline and CDP-ethanolamine Kennedy pathway [44, 45] (Figure 2). Both brunches of the pathway depend on the availability of the extracellular substrates, choline and ethanolamine, and on their entrance into the cells. Those substrates are polar molecules and have to be actively transported into the cells. The transport system for ethanolamine has not been extensively studied and remains poorly understood. Choline transport for PC synthesis is known to be mediated by a group of ubiquitous solute carriers of the SLC44A family, mainly by the member A1, also known as the choline transporter-like protein 1 (CTL1) and as the cell surface antigen CDw92 [46]. Immediately after entering the cells, choline and ethanolamine are phosphorylated by choline and ethanolamine kinases, of which

and - were identified and fully characterized in mammalian systems [47]. The kinase products, phosphocholine and phosphoethanolamine, are then coupled with CTP by the specific and the pathway regulatory enzymes CDP-phosphocholine and CDP phosphoethanolamine cytidylyltransferases (CCT/Pcyt1 and ECT/Pcyt2) to yield CDP-choline and CDP-ethanolamine, respectively, and to release inorganic pyrophosphate. In the final steps, the CDP-choline and the CDP-ethanolamine derivatives are condensed with diacylglycerol, catalysed by multiple DAG-choline and DAG-ethanolamine phosphotransferases (CPT, EPT, CEPT), to release CDP and to produce the bilayer forming phospholipids PC and PE at the endoplasmic reticulum (ER).


Pathways for the synthesis of phospholipids and their interconnection. The two branches of the Kennedy pathway are represented in grey. The key metabolites and enzymes catalysing the respective reactions are indicated. Abbreviations are as indicated in the main text.

The major regulatory points for the PC and PE synthesis through Kennedy pathway are the slowest (rate-limiting) reactions governed by Pcyt1 and Pcyt2, respectively. Also, as in other metabolic pathways, the Kennedy pathway is regulated by the substrate (choline/ethanolamine and DAG) availabilities. The regulation of Pcyt1 and Pcyt2 is extensively studied and has historically being considered similar, but evidence is emerging that they are completely distinct and differently regulated [48–53].

The rate of PC synthesis is predominantly regulated by Pcyt1 activity [54]. Pcyt1 is activated by association with membranes and downregulated by phosphorylation [55]. The translocation of Pcyt1 into cell membranes is stimulated by fatty acids as well as anionic phospholipids [56–59]. They induce the Pcyt1 binding to membranes, because of their negatively charged head groups [59]. Thus, changes in the lipid composition of the membranes can modify Pcyt1 activity, which typically leads to stimulation of PC synthesis.

Yet another regulatory step for PC synthesis is the choline availability which is regulated by the protein-mediated transport at the level of the plasma membrane. The universal transporter CTL1 transports choline in various tissues [60–62] and is considered predominant choline transporter in nonneuronal cells [46, 60, 62, 63]. Recently, it has been found that the CTL1 expression, choline uptake, and PC synthesis could be directly regulated by choline deficiency [64] and saturated and unsaturated fatty acids (FAs) in skeletal muscle cells [65]. The availability of choline and the type of FAs regulate PC synthesis together with differently modifying choline transport, PC synthesis, and DAG/TAG homeostasis. In choline deficient cells, PC synthesis from choline was blunted, which led to an increased availability of DAG and FA for TAG synthesis [64]. Similar to choline deficiency, chronic exposure of muscle cells to palmitic acid reduced CTL1, choline uptake, and PC synthesis and increased DAG and TAG [65]. In contrast, oleic acid showed no effect on CTL1 and choline uptake however, it increased PC synthesis at the level of Pcyt1 and elevated TAG content [65]. Oleic acid is a well-known stimulator of Pcyt1 membrane binding and activity [56–59].

The de novo synthesis of PE could be limited by the availability of substrates ethanolamine [66, 67] and DAG [68] or the enzymes EK [69] and Pcyt2 [53]. Based on the numerous radiolabeling studies and animal models, Pcyt2 is the main regulatory enzyme in the CDP-ethanolamine pathway. Heterozygous mice for the Pcyt2 gene (

) have reduced formation of CDP-ethanolamine, which limits the PE synthesis and increases the availability of DAG for TAG synthesis [70]. On the other hand, the overexpression of Pcyt2 could not accelerate PE synthesis when DAG availability for the last step in the pathway was limiting the PE formation [68], whereas EK overexpression accelerated PE synthesis in Cos-7 cells [69]. At low levels of ethanolamine, Pcyt2 limits the reaction rate, whereas at high levels of ethanolamine, EK limits the rate [71]. In ethanolamine deficient media, cells adapt the mitochondrial PS decarboxylation pathway to produce additional PE from serine (see Section 4) [72]. However, this is not the case in normal metabolism. Animal studies indicate that CDP-ethanolamine pathway is the major contributing pathway for the synthesis of PE [53].

In the liver, an alternative pathway utilizes PE to produce more PC and choline in a three-step methylation of PE by S-adenosylmethionine (SAM) catalysed by phosphatidylethanolamine-N-methyltransferase (PEMT). The PEMT pathway accounts for

30% of the hepatic PC synthesis and is coregulated with the CDP-choline Kennedy pathway [73]. It is not known how the CDP-ethanolamine Kennedy pathway, which only makes PE de novo, is linked with the PE used in the PEMT pathway for the formation of PC. However, in the PEMT knockout mice, the CDP-choline pathway is upregulated, whereas during the PEMT overexpression in hepatoma cells, it is downregulated, showing a strong link of the PEMT pathway with the liver PC metabolism [74, 75]. As a backup pathway, PEMT supplies PC for the very-low-density lipoprotein (VLDL) assembly and for the bile production and is a source of choline for the betaine synthesis in mitochondria [76]. The liver methylation of PE also provides PC for PS synthesis by the base-exchange reaction catalysed by PS synthase 1 (PSS1). The newly formed PS could be then transformed into mitochondrial PE by PS decarboxylase (PSD), forming a specific liver PE cycle [77] (Figure 2). The liver cycle PE-PC-PS may be involved in the maintenance of phospholipid levels when de novo CDP-choline and CDP-ethanolamine pathways are impaired.

In mammals, PS could be only made from the preexisting phospholipids (PC and PE) and L-serine, catalysed by PS synthases 1 and 2 (PSS1 and PSS2) in the mitochondria associated membranes (MAM), a subfraction of the ER. MAM have been proposed to be a distinct domain of the ER that comes into close contact with OMM and thereby mediates the import of newly synthesized PS into mitochondria to be decarboxylated into PE. The PS decarboxylation by PSD to form mitochondrial PE is a key function of PS in mitochondria (see Section 4).

The synthesis of PS is regulated by phosphorylation of PS synthase in yeast and mammalian cells [78, 79]. The major mechanism for regulating PS synthesis in mammalian cells is a feedback mechanism whereby the activity of PS synthases 1 and 2 is regulated by the end-product, PS [77, 80, 81]. In addition, PSS1 and PSS2 differentially modulate phospholipid metabolism. Overexpression of PSS1 in hepatoma cells decreases the rate of PE synthesis via the CDP-ethanolamine pathway [82], whereas overexpression of PSS2 does not [83].

4. The Biosynthesis of Mitochondrial Phospholipids

The maintenance of the mitochondrial bilayer and of a defined composition of mitochondrial phospholipids relies on the organelle capacity to synthesize CL, PE, PG, and PA in situ and on the external supply of PC and PS, which are exclusively synthesized in the ER and MAM and must be imported into the mitochondria [8].

PE is made in mitochondria by decarboxylation of PS. This reaction is catalysed by the inner mitochondrial enzyme phosphatidylserine decarboxylase (PSD). Even though PE produced by the CDP-ethanolamine pathway can be imported to IMM, the PS decarboxylation provides the majority of mitochondrial PE [84]. PSD knockout mice have mitochondrial dysfunction and die in the embryonic phase [25]. In addition, the mitochondrial PE deficiency resulting from siRNA silencing of PSD in CHO cells alters mitochondrial morphology and affects the ATP production, oxygen consumption, and the activity of the electron-transport components [85].

Cardiolipin, an important phospholipid in the mitochondrial membranes, is synthesized via condensation of phosphatidylglycerol (PG) and CDP-diacylglycerol catalysed by cardiolipin synthase (CLS) in the matrix side of the inner mitochondrial membrane (Figure 2). PG is produced by a two-step reaction: glycerol-3-phosphate and CDP-diacylglycerol condense to phosphatidylglycerol phosphate, which is then dephosphorylated to PG. Lack of CLS reduces the activity of the oxidative phosphorylation system and the import of proteins into mitochondria in yeast [86], but interestingly, the fundamental functions of mitochondria remain intact [87]. The regulation of CL synthesis was found to involve multiple mechanisms, such as factors that affect mitochondrial biogenesis, matrix PH, and respiration [88]. Recently, Tam41 (translocator and maintenance protein 41), a component of the mitochondrial translocator system, was found to regulate CL synthesis [23]. Tam41 mutant cells show an almost complete absence of CL and PG, suggesting that regulation may occur at the level of CDP-diacylglycerol (CDP-DAG) synthase. This study demonstrates that Tam41 is primarily required for PG and CL biosynthesis and that defects in protein import in Tam41 deficient cells are a consequence of the loss of PG and CL [23].

Finally, the phospholipid phosphatidic acid (PA) represents a branch-point for the synthesis of all phospholipids [89] (Figure 2). PA may be converted to CDP-DAG by the CDP-DAG synthase for the synthesis of PG, PS, and CL in a reaction catalysed by CTP:PA cytidylyltransferase [90]. Alternatively, PA may be converted to DAG, which is a substrate for the synthesis of PE and PC, in a reaction catalysed by PA phosphatases such as Lipin-1 [35, 91]. The PA phosphatases are one of the most highly regulated enzymes in lipid metabolism. Their activity is governed by phosphorylation, association with membranes, and modulation by components of lipid metabolism, such as CL and CDP-DAG [92, 93].

The synthesis of PA occurs by two acylation steps first glycerol-3-phosphate is converted to 1-acylglycerol-3-phosphate (lysophosphatidic acid) by glycerol-3-phosphate acyltransferase. The 1-acyl-sn-glycerol-3-phosphate product is then acylated to 1,2-diacyl-sn-glycerol-3-phosphate (phosphatidic acid) by 1-acyl-sn-glycerol-3-phosphate acyltransferase [94]. Studies have indicated that both lysophosphatidic acid and phosphatidic acid are synthesized on the outer surface of the mitochondrial outer membrane and then phosphatidic acid moves to the inner mitochondrial membrane where it serves as a precursor for cardiolipin biosynthesis [95]. PA can also be generated via phosphorylation of DAG through the action of a large family of DAG kinases (DAGK) or by hydrolysis of the phospholipids PC and CL by phospholipase D (see Section 8). In turn, PA can be metabolized to lysophosphatidic acid by phospholipase A2 or dephosphorylated to DAG by Lipin-1.

5. Trafficking of Phospholipids into and out of Mitochondria

Intracellular trafficking of phospholipids plays a crucial role in phospholipid homeostasis and provides phospholipids for cell and organelle membranes, including the OMM and IMM. Since most phospholipids, such as PE, PS, and PC, are synthesized in the ER, they have to be imported into the mitochondria. The import of PE produced by the CDP-ethanolamine pathway into IMM was found to be effective in CHO cells, HeLa cells, and yeast [96–98]. However, most of the PE in the mitochondria derives from the PS decarboxylation [85]. The synthesis of PE from PS in the IMM requires the translocation of PS from its synthesis sites in the ER. In addition, several studies have shown that PE formed from PS in the mitochondria could be exported out of this organelle [85, 97]. The transport of PS-derived PE out of the mitochondria was found to be affected by the rate of PE synthesis via the CDP-ethanolamine pathway and to be driven by a concentration gradient [97].

The mechanism for transport of phospholipids into and out of the mitochondrial membranes has been proposed to involve transient membrane contact sites. These contact sites occur in various organelles including the ER and mitochondria and may be involved in the trafficking of PS into and PE out of the mitochondria [99, 100]. Recent studies found that the ER membrane is physically tethered to the OMM by the ER-mitochondria encounter structure (ERMES) [101]. The ERMES is composed of a five-protein complex resident of both ER and mitochondria [101]. The main function of ERMES is to act as a mechanical link between the ER and mitochondria and to provide phospholipid exchange between these organelles [102, 103]. A defective ERMES complex alters phospholipid levels in the mitochondrial membranes and causes mitochondrial morphological defects [103, 104]. However, the flow of phospholipids between ER and mitochondria is not completely abolished in ERMES deficient cells, suggesting that additional ERMES-independent pathways for phospholipids transport must also exist [102].

Lipid transfer proteins, such as PC-transfer protein and nonspecific lipid transfer protein, have been shown to transfer phospholipids into the plasma membrane and may be also involved in the exchange of phospholipids between organelle membranes [105]. However, no transfer protein mediating PC, PS, and PE trafficking into/out of mitochondria has been established to date.

The rate of PS import to and PE export from the mitochondria is regulated by the acyl chain composition and by the metabolic rate [97]. Labeling studies showed that polyunsaturated PS species are preferentially decarboxylated, that is, imported to IMM [97]. In turn, PE derived from decarboxylation of PS contains the major polyunsaturated species as well as monounsaturated 36 : 1, whereas PE produced via the CDP-ethanolamine pathway contains several mono- and di-unsaturated species. The PE species composition of IMM and ER shows that most of PE in IMM derives from imported PS, while most of PE in ER is synthesized via the CDP-ethanolamine pathway [97]. In addition, Kainu and colleagues found that increasing the PE synthesis by CDP-ethanolamine pathway reduces the export of PS-derived PE from IMM. Similarly, the translocation of PS to mitochondria is coupled to its synthesis in the ER. However, the rate of transport of newly synthesized PS to mitochondria does not directly correlate with their rate of synthesis but depends on its hydrophobicity [97].

6. Intramitochondrial Trafficking of Phospholipids

The transbilayer movements between the mitochondrial leaflets must exist to allow the trafficking and accumulation of phospholipids, either when imported from ER or when synthesized at the IMM. Several studies have shown that the import of PS into mitochondria is mediated by the membrane contact between MAM, a subfraction of the ER, and OMM [84, 85, 97, 106, 107]. The PS must then translocate across the OMM and subsequently be delivered from the inner leaflet of the OMM, across the intermembrane space, to the outer leaflet of the IMM, which is the active site for PS decarboxylation to PE [108]. The mechanisms underlying the intramitochondrial lipid movement have been also suggested to occur via membrane contact sites [99, 109]. The model that best describes the intramitochondrial transport of PS involves pores in the OMM that lead to its movement from the outer leaflet of the OMM to the inner leaflet of the OMM. PS then diffuses further to the IMM along lipid bridges, which are analogous to the membrane contact sites, joining the two membranes [110]. Once in the IMM, PS can be converted to PE. The PE produced in the IMM can then diffuse back to the outer monolayer of OMM along a reverse route.

Phospholipid scramblases (PLS), members of the family of transmembrane lipid transporters known as flippases, are enzymes responsible for bidirectional movement of phospholipids between two compartments. Of this family, PLS3 was identified in mitochondria and it modulates translocation of CL from the IMM to the OMM, affecting mitochondrial structure, respiration, and apoptosis [111]. It is not known if PLS3 regulates transport of other phospholipids. The intermembrane space proteins Ups1 and Ups2 are responsible for CL and PE trafficking between the outer and inner mitochondrial membranes [112]. Loss of intramitochondrial CL trafficking in Ups1 deficient cells alters mitochondrial phospholipid composition and morphology, similar to ERMES deficient cells [112]. Mdm35, a common binding partner of Ups1 and Ups2 in the intermembrane space, also provides a coordinated regulation of PE and CL trafficking by these conserved regulatory proteins [113].

Therefore, the trafficking of phospholipids into mitochondria and intramitochondrial space provides this organelle with newly synthetized phospholipids, which are either essential for mitochondrial membrane composition or precursor for the synthesis of specific mitochondrial phospholipids. The balance in phospholipid trafficking, together with their synthesis and degradation, is essential for the maintenance of phospholipid homeostasis. However, not only the class of phospholipids accumulated in the mitochondrial membrane, but also the fatty acyl chain present within phospholipids is important for mitochondrial structure and function, which will be described in the next section.

7. Remodeling of Mitochondrial Phospholipids and the Role of Dietary Lipids

After the de novo synthesis of phospholipids, many of them undergo acyl chain remodeling, known as Lands’ cycle. The variation in chain length and degree of unsaturation of fatty acids present within phospholipids contributes to the diversity of mitochondrial membrane phospholipids, which is important for the biophysical properties of the membrane [114, 115]. In addition, activation of enzymes in the inner mitochondrial membrane requires acyl chain remodeling. For example, impairment of CL remodeling may interfere with assembly and stability of the respiratory chain proteins [115]. Conventionally, acyl chain remodeling involves the family of phospholipases A (PLAs), which catalyse the removal of an acyl chain from the glycerol moiety, and transacylases, which mediate reacylation with different FAs [114, 116].

Remodeling of cardiolipin has been extensively studied due to its role in the functionality of the mitochondria. CL remodeling involves tafazzin, a transacylase which generates specific patterns of CL species [117]. Deficiency of tafazzin alters CL composition and is related to dramatic changes in mitochondrial morphology and to Barth syndrome [118]. Barth syndrome is an X-linked recessive disease characterized by decreased levels of CL due to mutation in the tafazzin gene [119]. In Barth patients, a single molecular species, namely, tetralinoleoyl-cardiolipin, is missing, whereas other cardiolipins are either unaffected or even increased. This is caused by a reduced incorporation of linoleic acid (C18:2) into CL because of impaired FA remodeling by tafazzin [118, 120]. Changes in the cardiolipin FA composition in Barth syndrome may also interfere with assembly and stability of the oxidative phosphorylation complexes.

Another enzyme involved in cardiolipin remodeling is acyl coenzyme A thioesterase (Them5). Mice lacking Them5 have an altered mitochondrial morphology and function in hepatocytes [121]. Similarly, deficiency of lipocalin-2, a lipid transfer protein, alters the fatty acid composition of the PE, PC, and PS as well as the amount of mitochondrial cardiolipin in the mouse heart [122]. Lipocalin was found to reduce the linoleic acid (C18:2) content in phospholipids and to adversely affect mitochondrial function and energy production [122]. Indeed, cardiolipin enriched with symmetric linoleic acid allows for optimal function of the mitochondria [123].

The fatty acid composition of the diet can additionally modify the acyl chain remodeling of phospholipids and the mitochondrial function. In one study, mice exposure to rapeseed oil-rich diet showed altered mitochondrial membrane phospholipid composition in terms of both the type of acyl chains within the phospholipids and the proportions of phospholipid classes. As a result of altered mitochondrial membrane composition, these mice showed altered hepatic mitochondrial bioenergetics [124]. Guderley and collaborators showed that in trout mitochondria the changes in membrane phospholipid characteristics, including the acyl-chain composition, follow the patterns of fatty acids present in the diet. In this same study, the respiratory capacity of the red muscle mitochondria was altered by the different diets and was found to be higher in the diet rich in polyunsaturated lipids [125]. Several other studies in mammals have also demonstrated that dietary changes modify the FA composition of the major mitochondrial phospholipids [126–128] and, in particular, the molecular species associated with mitochondrial CL [57]. Following a diet deficient in linoleic acid (C18:2), an essential fatty acid, the rat heart showed a significant decrease in tetralinoleoyl CL, which affected mitochondrial oxygen consumption [127, 129]. Conversely, dietary supplementation with linoleic acid restored tetralinoleoyl CL in cultured fibroblasts from Barth syndrome patients and elevated CL levels [130].

The composition of mitochondrial CL was found to be altered by both quantity and quality of the dietary fat [131]. In hepatocytes, the composition of mitochondrial CL was altered in rats fed 30% fat diets in comparison to rats fed 5% fat diets. Also, the content of monounsaturated fatty acid (MUFA) and n-3 polyunsaturated fatty acids (PUFA) was increased with fish oil-rich diet in comparison to basal diet, both at 5% and 30% of fat in diet. Both total CL content and its C18:2 content were increased with liver steatosis and correlated to the activity of the ATP synthase [131]. Recently, a mitochondrial lipidomics analysis showed that the FA composition of the CL pool is not directly related to the presence of a given FA in the diet, but there is a selection for individual FA chains to be incorporated into the CL pool [132]. For example, the incorporation of 18:2 was found to be similar, despite diets varying

4-fold in this essential FA.

The n-3 PUFA content of the plasma membrane in different mouse tissues had the greatest sensitivity to changes in dietary fatty acids [133]. The fact that both n-6 and n-3 PUFA classes cannot be synthesised de novo by mammals suggests that the composition of membrane phospholipids may be strongly influenced by the ratio and abundance of n-6 and n-3 PUFAs in the diet. Several studies have shown that increasing the PUFA content of the diet increases the metabolic rate [134–136]. For example, treatment of rats with n-3 PUFA decreased proton leakage from the respiratory chain and this was related to the incorporation of PUFAs into mitochondrial PC, PE, and CL [137]. Treatment with docosahexaenoic acid (DHA), but not eicosapentaenoic acid (EPA), profoundly alters fatty acid composition of mitochondrial phospholipids in the rat heart, decreasing arachidonic acid and increasing DHA content. The increased DHA content delayed the mitochondrial permeability transition pore (MPTP) opening, which is associated with apoptosis and myocardial damage during ischemia [127]. DHA incorporation into CL was also found to regulate mitochondrial lipid-protein clustering, which alters several aspects of mitochondrial function [17]. Taken collectively, these results suggest that increasing the level of PUFA in the diet changes the composition of mitochondrial membrane phospholipids and consequently may alter mitochondrial capacity and function.

The response to dietary fatty acids also varies according to the phospholipid class. PC was found to be more responsive to variation in dietary MUFA content than the other phospholipid classes, whereas PE was more responsive than PC to both dietary n-6 PUFA and the n-3 PUFA [133]. Interestingly, Pcyt2 deficient mice, which have a decreased rate of PE synthesis, are PUFA deficient and have specifically modified FA composition in PE and in TAG [70]. Therefore, change in the phospholipid fatty acyl chain composition relies not only on the dietary fatty acid profile but also on the classes of phospholipids affected.

Taken together, these studies demonstrated that disturbance of enzymes involved in phospholipid remodeling, as well as changes in the dietary fatty acids, may alter the phospholipid composition of the mitochondrial membrane. Dietary interventions that are able to influence mitochondrial membrane phospholipids, hence modifying its physical properties, respiration, and other processes such as MPTP, are emerging as novel therapeutic strategies [124]. Therapies addressed to mitochondrial phospholipids have been suggested as potentially useful to treat pathologies, such as cancer, cardiovascular and neurodegenerative diseases, obesity, and metabolic disorders [124].

8. Degradation of Mitochondrial Phospholipids

Many phospholipids have a rapid turnover, indicating that their degradation plays an important role in membrane homeostasis. The phospholipid degradation is mainly catalysed by nonlysosomal phospholipases. These phospholipases are divided into three classes based on the bound they cleave: phospholipases A (PLAs), phospholipases C (PLCs), and phospholipases D (PLD).

PLAs release the fatty acid in the sn-1 and sn-2 position of the glycerol moiety generating lysophospholipid and free FA. The lysophospholipid is reacylated to generate new phospholipid, the mechanism known as FA remodeling, or it is degraded by a lysophospholipase. Several studies suggest that PLAs mediate much of the phospholipids turnover [138, 139]. The PLA2 family, which hydrolyses FA bound at the sn-2 position of phospholipid, has been related to turnover and remodeling of phospholipids as well as to membrane homeostasis [140, 141]. iPLA2

-independent PLA2 family, is preferentially distributed in the mitochondria. Because of the association of iPLA2 with mitochondrial membranes, this enzyme may be involved in integrating phospholipid and energy metabolism. Mice null for iPLA2 display abnormal mitochondrial function with a dramatic decrease in oxygen consumption [142]. Thus, iPLA2 is essential for maintaining bioenergetic mitochondrial function through regulation of mitochondrial membrane phospholipid homeostasis. Because, its role in cardiolipin remodeling, the hippocampus of iPLA2 null mice shows an elevated content of mitochondrial cardiolipin with an altered chain length composition [142].

Phospholipases C (PLCs) hydrolyse the bond between glycerol backbone and phosphate to yield DAG and phosphorylated head group. PLCs are generally involved in generation of second messengers for signalling events [143], but they also play a role in the turnover of PC and PE [144, 145].

Phospholipases D hydrolyse the bond between phosphate and the head group to yield PA and free head group. Both classical phospholipase D (PLD) family members found on many cytoplasmic membrane surfaces and MitoPLD, the PLD family member found on the mitochondrial surface, can generate PA via hydrolysis of the phospholipids PC and CL [26]. MitoPLD-generated PA regulates mitochondrial shape through facilitating mitochondrial fusion [146]. Mitochondrial fusion and fission are very important in maintaining mitochondrial and cellular function, and morphological changes of the mitochondria are linked to cell apoptosis and neurodegenerative disease [146]. For example, in brains of Alzheimer’s disease patients, PLD1 is upregulated in the mitochondrial membrane, which affects the composition of mitochondrial membrane phospholipids [147].

The involvement of phospholipases in phospholipid homeostasis is also demonstrated by PLD-like enzymes that catalyse PS synthesis by base-exchange reactions. PS synthases 1 and 2 (PSS1/2), the PLD-like enzymes, mainly catalyse the exchange of the head group of PE or PC for serine rather than the exchange of the serine head group of PS for choline or ethanolamine. Thus, PLD and PLD-like enzymes could play an important role in different phospholipid homeostasis [84].

9. Conclusion

The identification of a link between phospholipids and proteins in the mitochondrial membrane has enhanced our understanding of mitochondrial morphology and function. The regulation of synthesis, trafficking, and degradation of phospholipids is essential to maintain phospholipid homeostasis in the mitochondria. Remodeling of phospholipid in the mitochondria can be modified by dietary fatty acids, which contribute to mitochondrial membrane integrity. However, the molecular mechanism for the coordination of synthesis, degradation, trafficking, and remodeling of phospholipids is still an active area of research. For example, how is the synthesis and degradation interconnected? Which are the signals for phospholipid transport into and out of mitochondria? How can diet be used to modulate phospholipid homeostasis in health and disease states? Undoubtedly, many discoveries will be achieved in a few years.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.

Acknowledgments

This work in the laboratory of Marica Bakovic was supported by the Canadian Institute of Health Research and the National Sciences and Engineering Research Council of Canada.

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Copyright

Copyright © 2014 Laila Cigana Schenkel and Marica Bakovic. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


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Discussion

Caspase-8 interacts with mitochondria in both healthy [52] and apoptotic [53], [54] cells. However, it has remained unclear how caspase-8 interacts with CL in mitochondria. It has been suggested that this enzyme is translocated into mitochondria together with its known substrate, Bid [25]. However, caspase-8 translocation to the mitochondria after Fas activation is unaffected in Bid knock-down cells. Caspase-8 interaction with mitochondria may be mediated by other proteins [54] or, as described for tBid, caspase-8 may interact not only with other proteins, but also directly with the lipid CL at the mitochondrial membrane. The 𠇎mbedded together” model for the association of Bcl-2 family members with the lipid domain of membranes assumes that the insertion of these proteins into the mitochondrial outer membrane during apoptosis affects the affinities of the various Bcl-2 proteins, creating new interaction surfaces [30], [55]. It has been conjectured that mitochondrial-membrane microdomains enriched in CL play an important role in apoptosis and enzyme flux control [56].

We investigated the role of CL in the formation of such an apoptosis-activating reaction platform, by generating a minimal in vitro reconstitution system with biomimetic membranes (LUVs and GUVs). Western blotting and flow cytometry ( Fig. 1 ) were used to distinguish between the specific and non-specific binding of Bid and caspase-8. Indeed, whereas Bid interacted with neither DOPC-only nor CL + -liposomes, caspase-8 was found to interact with CL-containing LUVs, giving rise to the p43 kDa CL-activated form ( Fig. 1b, c ).

We then used Laurdan as a fluidity tracer, to study the effects of caspase-8, Bid and the caspase-8+ Bid complex on a relevant membrane model. Differences in the excitation and emission fluorescence spectra of Laurdan in the gel and liquid-crystalline phase make it possible to use the general polarization (GP) parameter to report on the local changes in membrane water content related to changes in membrane fluidity due to protein binding. Bid alone did not bind to liposomes ( Fig. 4 ). By contrast, caspase-8 and caspase-8 plus Bid decreased the fluidity of CL-containing membranes ( Fig. 2 ), as, to a lesser extent, did tBid. This result is consistent with previous data indicating that the presence of tBid may promote the formation of highly curved non-lamellar phases [57]. One surprising finding was the marked effect of procaspase-8 itself on the membrane and subsequent Bid binding. The additive effect of procaspase-8 and Bid may result from the acquisition of full functional activity upon binding to CL. The interaction of CL with caspase-8 on the membrane is important for the progression of apoptosis, with the formation of a local CL-protein reaction platform evident from the change in GP value. These results shed light on the role of mitochondrial membranes in the regulation of Bcl-2 protein family activity [33].

The results of rupture-tension experiments and those for Laurdan fluorescence are complementary. The rupture-tension approach, originally developed by Evans and coworkers [see [43] and citations therein], can be used to quantify the micro-mechanical properties of a thin film, such as a lipid membrane, by studying its deformation. Here, the expansion modulus (Ks) and the rupture tension (τr) were evaluated by expanding large unilamellar vesicles (GUVs), the membranes of which constituted a model system mimicking mitochondrial contact sites. We found that the addition of CL resulted in a marked decrease in the elastic moduli of DOPC lipid bilayers, with both Ks and τr strongly affected ( Fig. 3b and 3c ). The decrease in Ks following the addition of CL indicates that the membrane becomes easier to expand in the presence of this lipid ( Fig. 3b ). The CL molecule has an inherent conical shape in a pure phase system, it would therefore preferentially be found in the inverted hexagonal phase. In the model system used here (CL/DOPC =𠂥/95 mol/mol), we intentionally avoided setting up such a condition: Each CL molecule was surrounded by DOPC molecules. In theory, the system displayed almost ideal mixing, as the chains of the two lipids (oleoyl-CL and DOPC) were identical. The predominance of the species preferring a lamellar phase ensured the maintenance of a lamellar state. The CL was, thus, structurally frustrated as it was embedded as a minor component within its host membrane. Nevertheless, its presence locally modifies spontaneous curvature. Due to their four hydrocarbon chains, CL molecules subjected to external force act like integrated springs that can be expanded more easily than the DOPC molecules, resulting in a lower Ks. However, it is not possible for the system to assume a hexagonal phase and the limits of expansion of the lamellar phase are soon reached. The rupture tension is, therefore, lower than that for the pure system. The data for the pure control system are consistent with published data for DOPC vesicles [58], giving a Ks value of about 200 mN/m.

We then assessed the mechanical consequences of proteins in the pure control system and in the PC/CL contact site model ( Fig. 3 ). None of the proteins tested interacted readily with the pure DOPC control membrane. The properties of CL-containing GUVs were not changed by Bid binding, whereas the binding of caspase-8, tBid and caspase-8 plus Bid clearly modified the mechanical properties of these vesicles ( Fig. 3b and 3c ). The binding of caspase-8 alone partly reversed the effects of CL, indicating a role for CL in binding. The structural frustration observed when CL alone is added was reduced, such that the expansion module value was between those for the control and the DOPC/CL model system. The tensile breaking strength was essentially the same as that for the pure system, being limited only by the lipid membrane itself. Most probably, caspase-8 detects the curvature frustration close to CL locations within the membrane, and its insertion partially compensates for it. tBid alone also bound to the model vesicles (DOPC/CL). In this case, the expansive elastic response of the membrane, assessed by calculating the modulus Ks, was fully restored to that of the pure DOPC control system: The adsorption of this protein fully released the structural frustration caused by the presence of CL. It is likely that all of the interaction sites were saturated. Nevertheless, the presence of the protein clearly caused defects that weakened the membrane to mechanical stress. This is evident from the very low value of the rupture tension. Although the membrane initially responded to a deformation force with an increase in area similar to that for the pure system, the total range of expandability was much lower, and the membrane broke down when the tension increased by ∼ 4.2 mN/m, corresponding to a change of ∼ 70% with respect to the control systems (pure DOPC or DOPC/caspase-8). Evidently, two domains with different elastic properties were formed. A major part of the membrane consists of essentially pure DOPC and does not participate in the interaction, or establishment of a reaction platform. Its elastic properties are therefore not modified, such that the observed Ks was � mN/m. The other part of the membrane, which contains CL as the initiator of a reaction platform, is more rigid. It does not discernibly contribute to membrane expansability but it limits the overall strength, as shown by the low value of τr. A similar behaviour was observed for caspase-8 plus Bid, within the limits of experimental resolution, and in line with the GP results obtained with LUVs. All these findings are consistent with the recently described interactions between Bcl-XL [59] and tBid. We confirmed that CL plays an essential role in the association between caspase-8 and biomimetic membranes ( Fig. 6 ), and most probably also biological membranes [25]. We suggest that CL is a component of the reaction platform formed subsequently (which also contain caspase-8 and Bid), in which it acts as a cofactor for caspase-8 activation. As the platform is formed, it immediately acquires its enzymatic function but only if CL is present ( Fig. 4 and Fig. 5 ). The production of tBid in the presence of caspase-8, when it interacts which CL, promotes vesicle breakdown this effect is inhibited if caspase-8 inhibitors are added to the system [41]. These results indicate that the presence of caspase-8 linked to CL is essential for the formation of the so-called “mitosome” [25], [41]. In addition to interactions between CL and caspase-8, there may also be protein-protein interactions in vivo. It remains unclear whether other proteins, such as Rab5 [60], [61], which requires caspase-8 activation, or BAR [54] and FLASH, which mediate caspase-8 translocation to mitochondria [62], [63], [64], play an auxiliary role in the functional relationship between caspase-8 and CL. Possibly, MTCH2/MIMP [65] and its role in tBid recruitment may act in synergy with CL-induced mitosome formation to facilitate MOMP. The work we report here expands our knowledge of Bid-induced pro-apoptotic signalling and provides a description of the role of CL in capsase-8 recruitment and activation at the surface of the mitochondrial outer membrane. We are however far from grasping all the intricate and complex molecular alterations and interactions that lead to the activation of Bid, mitochondrial membrane permeabilisation and apoptosis via the mitochondrial pathway following stimulation of the death receptors.

(a) The diagram depicts the sequence of events in cells of type II according to Gonzalvez et al. [25]. The CL (red heads)/caspase-8 platform at the contact sites between inner and outer mitochondrial membranes (enriched in CL) binds BID resulting in the production of the active truncated, C-termimal part of BID (tcBID). This in turn causes CL induced perturbations of the membrane curvature, BAK/BAX oligomerization and cytochrome c release. (b) Schematic representation of the reconstituted functional platform on giant unilamellar vesicles containing CL with the p18/p10. DD, death domain DED, death effector domain p10 and p18 form the catalytic core of the caspase. The p43/p10 caspase-8 isoform comprises two DEDs, one p10 domain and one p18 domain. IMM, inner mitochondrial membrane IMS, inter membrane space OM, outer mitochondrial membrane. Red dots in the intermembrane espace, cytochrome c and the violet head correspond to the cardiolipin at the contact sites between outer and inner membrane.

The results that we present demonstrate and describe essential roles played by lipids in biological processes. In particular, they provide new insights into how mitochondrial specific lipids like CL can have active functions that go far beyond simply constituting a matrix for protein activities. Indeed, functional lipids appear to contribute not only to modulating the interactions between Bcl-2 family members, but also as key players in recognition processes as demonstrated by the example cardiolipin triggering the activation of caspase-8 in the apoptotic process.


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