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C2: Lipid Distribution in Cells - Biology

C2: Lipid Distribution in Cells - Biology



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To understand movement of lipids in an actual cell, a better understanding of lipid synthesis and trafficking in cells is important. et al) while the following figure shows how the lipids composition of membranes organelle membranes.

Table: Distribution of Lipids in Resting Macrophage

Lipid Categories

Nucleus

Mitochon-dria

Endo. Reti.

Plasma Memb

microsome

cytosol

Whole cell

Glycero-phospholipids

149

152

150

151

142

109

155

Prenol lipids

5

5

5

5

5

5

5

Sphingolipids

48

47

48

48

48

47

48

Sterol lipids

13

12

12

13

11

5

12

Total

215

216

215

217

206

166

220

Figure: Approximate Distribution Phospholipids in Resting Mammalian Cells

Lipids in membranes are often distributed asymmetrically. The inner and outer leaflet of a biological membrane usually have different PL compositions. For example, in red blood cell membranes, the outer leaflet is composed mostly of sphingomylein (SM) and PC, while the inner leaflet is composed mostly of PE and phosphatidyl serine (PS). This phospholipid contains the amino acid serine linked through its side chain (-CH2OH) to phosphate in position 3 of diacylglyerol. With a negative charge on the phosphate and carboxylate and a positive charge on the amine of PS, this phospholipid is acidic with a net negative charge. All the PS is located in the inner leaftet! This observation will become important latter on, when we discuss programmed cell death. A dying cell will expose PS in the outer leaflet. This is in fact one of the markers of a dying cell.

The membrane lipid composition in an average mammalian cell

Lipid%
PC45-55
PE15-25
PI10-15
PS5-10
PA1-2
SM5-10
cardiolipin (bis-PG)2-5
cholesterol10-20

Intermediate Filament Associated Proteins

Günther A. Rezniczek , . Gerhard Wiche , in Methods in Enzymology , 2016

4.7.2.2 Coimmunoprecipitation from Cell Fractions

Perform cell fractionation as described in Section 4.2.1 . Use the cytosolic and membrane fractions for coimmunoprecipitation. Before adding primary antibodies, an optional preclearing step can be performed. Add primary antibodies and incubate 3 h to overnight with end-over-end rotation. Add protein A (or G) sepharose and incubate for 3 h to overnight (incubation times need to be individually balanced between specificity and efficiency of precipitation). Remove the beads by careful centrifugation (see above) and wash 3–4 × with the respective lysis solution (inhibitors are not necessary).


Background

The ability to produce phosphatidylserine (PtdSer) is essential for mammalian survival [1], while the lack of PtdSer production in yeast leads to growth defects and an increase in other negatively charged lipids in an attempt at compensation [2, 3]. In addition, over production of PtdSer leads to the congenital disease Lenz-Majewski syndrome, characterized by the combination of sclerosing bone dysplasia, intellectual disability and distinct craniofacial, dental, cutaneous and distal-limb anomalies [4].

PtdSer has important roles in apoptosis and blood clotting, and most of what is known about PtdSer applies to these roles. However, in homeostasis PtdSer is not generally externally exposed, yet it clearly plays a vital role in healthy cells. The function of PtdSer, as with all lipids, is determined by both its concentration and sidedness in individual organellar membranes. Mitochondria associated membranes (MAMs) of the endoplasmic reticulum (ER) have high rates of PtdSer synthesis and serve as a conduit for the transfer of lipids between the ER and adjacent mitochondria [5, 6]. The bulk subcellular distribution of PtdSer results from the coordinated actions of metabolic enzymes in conjunction with vesicular and nonvesicular transport pathways, while the topology of PtdSer results from the actions of transmembrane enzymes capable of moving PtdSer between lipid bilayers PtdSer flippases, floppases, and scramblases [7, 8]. Until relatively recently, PtdSer distribution and topology studies depended solely on the fractionation and subsequent chemical analysis of cellular organelles. These early studies highlighted PtdSer distribution throughout the cell is unbalanced (Fig. 1a), being more concentrated in the plasma membrane (PM) (

10–15% total lipid) with lower levels in the ER (

1%), the latter of which uses PtdSer as a source of phosphatidylethanolamine (PtdEtn) (reviewed in [7, 9, 10]). The PtdSer content of less abundant organelles, including the endosomal system, has generally been less well defined because of the difficulty inherent in purifying them to homogeneity.

Intracellular distribution of PtdSer. a Relative abundance of PtdSer in membranes as mol% of total lipids throughout organelles of the cell. ER – endoplasmic reticulum, PM – plasma membrane. b, c The probe LactC2 labels cytoplasmic-facing leaflets containing PtdSer. When co-expressed with additional organellar markers (such as the plasma membrane labelling PH-PLC (b)) relative correlations as determined by calculation of Pearson’s correlative co-localization (c) can be determined as a proxy for the relative amounts of PtdSer in the cytoplasmic-facing leaflets of organelles (as first published in Hirama et al. [48]). Markers for plasma membrane (PH-PLC), ER (Sec61), Golgi (GalT), mitochondria (Mito (MitoTracker)), early endosomes (Rab5), fast and slow recycling endosomes (Rab4 and Rab11, respectively) and lysosome (LAMP1) are shown. The lack of ER and Golgi labeling by LactC2 suggests a lack of PtdSer in the cytoplasmic leaflets as discussed in the text

In addition to difference of PtdSer content amongst organelles, the unequal bilayer distribution of PtdSer at the PM has long been appreciated [11], as has the importance of movement of PtdSer from the cytoplasmic to exofacial face of the PM being involved in critical signaling events including blood clotting [12] and apoptotic cell recognition and removal by macrophages [13]. Furthermore, the PM has a net-negative charge on its cytoplasmic face [14], and consequently has an essential role in charge-based signaling events [15]. However, the contribution by PtdSer to this charge, as well as precise localization and the dynamics of PtdSer, or indeed other organelles, within whole and live cells, remains an area of active research which has recently been aided by new tools for the detection and visualization of PtdSer. In this review, we will highlight recent contributions to the understanding of PtdSer distribution and its roles within a normal cell.

Distribution and dynamics of phosphatidylserine

The development of the PtdSer-specific LactC2 probe, based on the PtdSer-specific calcium independent discoidin-type C2 binding domain of lactadherin (also known as Milk fat globule-EGF factor 8 (MFGE8)) [16] has enabled the visualization of PtdSer in live cells (Fig. 1b-c). Indeed, the initial study using this probe showed for the first time the cytoplasmic-facing distribution of PtdSer in live cells. This initial LactC2 study underscored the importance of PtdSer in providing the negative charge of the PM, finding that cationic probes track the presence of LactC2-identified PtdSer, including in the absence of polyphosphoinositides [16]. The study also highlighted the presence of PtdSer in, and its ability to recruit charge-based protein probes to, endosomal compartments, while not being detectable in the cytoplasmic-facing cis-Golgi, ER or mitochondria. While it is possible the LactC2 probe does not have high enough sensitivity to detect the relatively low levels of PtdSer present in these organelles [9, 10], it is also possible that, like in the PM, PtdSer leaflet distribution in intracellular organelle membranes is asymmetrical [17]. Indeed, there existed significant evidence prior to the development of the LactC2 probe suggesting this is the case, at least in the ER [18,19,20,21]. This evidence has since been strengthened with additional data that does not require the biochemical isolation, and potential disruption of, this intricate tubular organelle. Using a combined light microscopy and on-section staining electron microscopy (EM) approach, the LactC2 probe was able to detect PtdSer on the luminal but not cytoplasmic facing ER membrane [22]. A modified ER-targeted LactC2 probe has also been used to successfully detect PtdSer in the ER lumen of live cells [23].

The ability of PtdSer to change membrane leaflets faces a high energy barrier, with spontaneous translocation estimated to only occur in the order of hours per single molecular translocation event [24, 25]. Three categories of proteins have been characterized that enable the trans-leaflet movement of lipids: flippases that transfer lipids to the cytosolic leaflet from the PM extracellular or organellar luminal leaflet, floppases that transfer in the opposite direction (out of the cytosolic facing leaflet), and scramblases that are bidirectional [26,27,28]. As the cytoplasmic leaflet of the ER is where the active site of glycerophospholipid enzymes reside [29], it has generally been thought that most glycerophospholipids in the ER are scrambled equally between leaflets to allow for proper ER membrane expansion and leaflet coupling [30, 31]. How this can be compliant with PtdSer having a polarized distribution in the lumen of the ER is unclear. However, expression of gain-of-function PtdSer synthase 1 identified from Lenz-Majewski syndrome patients does result in the appearance of cytosolic PtdSer in the ER, demonstrating that the normal mechanism(s) that restrict PtdSer to the luminal leaflet are saturable [32]. One possibility is that PtdSer, once in the luminal leaflet, is kept there through interactions with luminal proteins and/or Ca 2+ [33]. Other non-mutually exclusive possibilities are that movement PtdSer from the cytoplasmic-facing leaflet occurs at the MAM into the mitochondria where it is used for the production of PtdEth [34], or PtdSer is removed from the cytoplasmic leaflet through non-vesicular transport by lipid transfer proteins (LTPs).

LTPs, along with vesicular trafficking, are how lipids move between cellular membranes [9, 33, 35]. Recent studies have highlighted the ability of specific LTPs, oxysterol-binding homology (Osh) proteins 6 and 7 in yeast [36, 37] and oxysterol-binding protein (OSBP)-related proteins (ORPs) 5 and 8 in mammalian cells [36, 38], to move PtdSer between membranes. The existence of these PtdSer-specific LTPs thus provide a potential mechanism for the generation and/or maintenance of the PtdSer cellular membrane gradient present in cells. Indeed, recent studies have shown that LTP-mediated transfer of PtdSer against its concentration gradient is possible through exchange with phosphatidylinositol 4-phosphate (PtdIns4P) down its concentration gradient from the PM to the ER, where the phosphatase Sac1 converts PtdIns4P to PtdIns [38, 39]. However, recent evidence suggests this exchange may be principally used to fine tune the PM levels of PtdIns4P and PtdIns(4,5)P2 rather than be responsible for bulk movement of PtdSer into the PM [39, 40]. There is also compelling evidence for the importance of vesicular trafficking in being the major route for PtdSer trafficking and concentration within the PM. For example, in yeast with temperature-sensitive mutations in secretory proteins Sec6 and Sec1, the polarization of PtdSer in the PM normally seen at a forming bud is inhibited and PtdSer instead accumulates on the vesicle that are prevented from fusing with the PM [2]. Additionally, endosomal recycling is important in the maintenance of high PtdSer levels, with inhibition causing a redistribution of PtdSer throughout the endosomal system in yeast [41]. Similarly, disrupting LTP function in mammalian cells has been found to result in slightly altered, but not disrupted, cellular membrane PtdSer distribution [38, 39]. Furthermore, Snx4, a member of the sorting nexin family of proteins involved in endosomal cargo sorting and recycling [42] that is specifically involved in recycling of Snc1 in yeast [43] and transferrin receptor in mammalian cells [44] has recently been implicated in leading to the modification of endosomal PtdSer levels [41].

Thus, while nonvesicular lipid transport, mediated by LTPs, play an important role, vesicular trafficking appears to be a significant contributor for maintaining the inter-membrane PtdSer gradient within the cell. Though the full molecular mechanisms of how PtdSer is segregated from other lipids remains to be fully elucidated, biochemical studies indicate a significant fraction of PtdSer in mammalian cells is enriched in PM-derived detergent-resistant, cholesterol-enriched “lipid-rafts” [45]. This biochemical data is supported by both electron microscopy analysis showing PtdSer is not homogenously distributed throughout the PM [22] and the finding that cholesterol and PtdSer co-segregate throughout subcellular compartments, being most concentrated in the PM and early endosomal compartments and relatively absent from the ER [22, 46, 47]. Further, acute changes in either affect the distribution of the other cholesterol is required for the normal distribution of PtdSer [2, 48] and acute changes in PM levels of PtdSer alter the distribution of cholesterol [46]. Evidence is also building for the likelihood that plasma membrane outer leaflet rafts, dependent on glycersphingolipids and cholesterol [49], are coupled to inner leaflet rafts [50, 51]. The importance of PtdSer in this coupling, in both the PM and endosomal membranes, is the subject of a recent excellent review [52] so will not be further covered here.

Roles of intracellular phosphatidylserine

As described in Background, PtdSer is essential in mammalian cells [1], while yeast lacking PtdSer are viable but have greatly reduced growth kinetics [2, 3]. As well, as PtdSer-mediated extracellular signaling, such as during blood clotting and apoptosis, has recently been reviewed [53,54,55], we will focus here on information regarding the roles of PtdSer within healthy non-apoptotic cells (Fig. 2).

Current knowledge of roles and intracellular transport of PtdSer. PtdSer is produced in the ER, from where it is distributed throughout the cell. PtdSer can be transferred to the mitochondria through mitochondria associated membranes (MAMs) (1), where it is mostly converted to PtdEtn. Distribution to the PM and endosomal system can occur via traditional vesicle-mediated trafficking as well as via direct movement via PtdSer-specific lipid transfer proteins (2). The relative importance of both trafficking methods is currently unclear. At the PM (3), PtdSer is kept in the cytoplasmic-facing leaflet and is important for generating a high net-negative charge. A number of important signaling molecules are recruited to the PM through charge and/or direct PtdSer recognition binding, with PtdSer thus playing essential roles in many signaling cascades and protein localization. PtdSer also plays important roles in endocytosis (4), including through its curvature-inducing headgroup interactions as well as interactions with proteins required for caveolae formation. PtdSer may also play a role in Golgi function (5), related to cargo sorting and budding from the trans-Golgi. PtdSer also appears to be important for recycling of cargo and interaction with the recycling machinery (e.g. Evectin2, EHD1, Snx4) at the recycling endosome (6). These interactions with the recycling machinery also likely helps to ensure PtdSer returns to, and maintains its enrichment on, the PM while causing reduced PtdSer levels on the late endosomes and lysosomes. Mito – mitochondria, ER – endoplasmic reticulum, PM – plasma membrane, EV – exocytic vesicle, EE – endocytic vesicle, RE – recycling endosome, Lys – lysosome

As described, at steady state in a healthy cell PtdSer makes up to

15 mol% of the total lipid in the PM. Furthermore, as it is nearly exclusively in the inner (cytoplasmic-facing) leaflet it can therefore make up to

30 mol% of the lipid on this leaflet. As the major lipid with a net-negative charge, PtdSer is therefore responsible for providing much of the inner leaflet’s charge density. A significant role of PtdSer then is interacting with proteins in a non-specific charge-based manner to permit their appropriate localization within the cell (Table 1). For example, the protein kinase Src and Ras GTPase family members Rac1 and K-Ras are proteins whose membrane targeting requires a polycationic stretch in addition to lipid modifications [56, 57]. The polycationic stretch of K-Ras4B has a net charge of + 8, resulting in its localization almost exclusively at the PM. If PtdSer is removed [58], or if the net charge of this stretch is varied the resulting mutants are directed additionally to other membranes constructs of intermediate charge (e.g., + 5) localize to endosomal membranes [16]. Similarly, Src has a polycationic stretch next to its myristoylated residue at the N-terminus with a net charge of + 5, and the kinase was found to associate not only with the PM but also extensively with PtdSer-enriched endosomal membranes [16].

Further evidence of the importance for PtdSer in charge-based protein distributions has been observed with the phagocytic process. When pathogens cause a depletion of PtdSer from phagosomes, Src is also lost [59]. In other instances, such charged motifs are not sufficient to direct proteins to a membrane but nonetheless influence their targeting, likely playing a complementary role [56, 60]. Evidence that this is the case comes from studies in yeast where polarized PtdSer is required for the recruitment of the signaling and polarity-regulating molecule Cdc42 to the forming bud neck without PtdSer Cdc42 remains Golgi-associated and buds are very inefficiently formed, leading to poor growth [2]. Similarly, Cdc42 and Rho1 are dependent on PtdSer polarization for their proper localization and function in Schizosaccharomyces pombe [61]. In yet another example, the plant GTPase Rho of Plants (ROP) family member ROP6 doesn’t appear to require PtdSer for its PM association, but does require PtdSer to be stabilized into nanodomains within the membrane upon activation that allows proper signal transduction [62]. Whether PtdSer is required for, or can modulate, signaling of other ROP family members, all of which contain a polybasic stretch of amino acids at their C-terminus [62], remains to be seen.

Traditionally, the interactions between polycationic stretches in proteins and anionic phospholipid headgroups have been thought to be strictly charge based with little specificity. However, recent evidence challenges this assumption. For instance, K-Ras4B which contains six lysine residues adjacent to a farnesylated cysteine residue, has recently been shown to interact with PtdSer preferentially [63]. The tail region of K-Ras4B adopts a series on conformations, disordered, ordered and intermediate, with the disordered being the preferred conformation. This conformation is also able to H-bond PtdSer more effectively than the other two confirmations [63]. Conversely, other proteins such as K-RasG12V and Rac1 show no preference for PtdSer [63,64,65]. While these are only initial studies, the results suggest that some polybasic proteins may have a preference for PtdSer or other anionic lipids beyond simple electrostatically driven interactions.

There are also multiple lines of evidence indicating the charge of PtdSer contributes to PM curvature and is important for the formation of some forms of endocytic vesicles. For example, caveolae are bulb-shaped nanodomains (50–100 nm) of the PM that have been linked with many physiological functions, including mechanosensing and endocytic transport [66]. While caveolae have been known to be enriched for cholesterol and specific glycerosphingolipids, including GM3 [67], PtdSer has recently been identified as being required for their formation and maintenance [68]. This is likely at least partly due to the charge-based PtdSer binding of the cavin1 protein [69] which, along with caveolin1, is required for in vivo caveola formation [70]. PtdSer is also capable of causing membrane curvature and induce endocytosis upon the acute removal of cholesterol, again a consequence of the charged headgroup of PtdSer [48]. It is likely that cholesterol, which makes up

40 mol% of PM lipids [10], helps to keep the PtdSer headgroup charge density on the inner leaflet low enough to not induce spontaneous curvature. However, once cholesterol is removed the distance between phospholipid headgroups is decreased, resulting in high spontaneous curvature capable of forming endocytic tubules [48, 71]. Indeed, increasing PtdSer levels on the inner leaflet of the PM above homeostatic levels (and therefore charge density) without concomitant cholesterol removal is also sufficient to increase formation of endocytic vesicles [48]. It is tempting to speculate that the cavin and caveolin proteins are taking advantage of this curvature-inducing property of PtdSer to induce caveolae. Thus, while cholesterol appears important for PtdSer cellular localization, it also appears to be important for modulation of PtdSer spacing and membrane curvature induction. This intimate relationship with cholesterol likely plays important roles in other PtdSer function as well, as suggested by PtdSer dynamics and interactions with caveolae [68] and signaling proteins [2, 59, 62].

The understanding of the role of PtdSer in internal membranes remains even less clear than the roles at the PM. Similar to the plasma membrane, recycling endosomes are rich in PtdSer [72] and recent work has demonstrated that PtdSer supports a variety of functions in these endosomes. The endosomal protein Evectin-2 contains a pleckstrin homology domain that binds to PtdSer rather than phosphoinositides [72]. Depletion of Evectin-2 or decreasing the availability of PtdSer prevents the movement of cholera toxin from the recycling endosome to the Golgi. Similarly, depletion of Evectin-2 and a reduction of PtdSer levels results in an inability of Golgi proteins (e.g. TGN38) to be retrieved from endosomes [72, 73]. In addition to the presence of PtdSer on the cytosolic leaflet of recycling endosomes, PtdSer flippases (e.g., ATP8A1, ATP8A2) are also required to support trafficking events. One critical effector downstream of flipped PtdSer is the Eps15 homology domain-containing protein-1 (EHD1), an ATPase with dynamin-like activity and a role in membrane remodeling required for the retrograde transport of Shiga toxin to the Golgi [74, 75]. Curiously, PtdSer, Evectin-2 and ATP8A1 have all recently been implicated as regulators of Yes-associated protein (YAP) signaling and cell proliferation [76]. ATP8A1 knockdown results in the activation of Lats, which in turn phosphorylates YAP and prevents its translocation into the nucleus. Silencing of Evectin-2 results in a decrease of Nedd4-mediated ubiquitination of Lats1, resulting in increased levels that also result in increased phosphorylation and inactivation of YAP. These studies raise several questions regarding how PtdSer and its flipping in recycling endosomes are controlling these effectors. Additionally, since recycling endosomes receive a lot of incoming membrane from the asymmetric plasma membrane, it is unclear where the luminal leaflet PtdSer is coming from to serve as a substrate for the flippases. Much is still to be learned regarding the cell physiology of PtdSer and we anticipate that the same biophysical properties PtdSer imposes on the plasma membrane will hold in endosomes and the trans-Golgi.


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Randomly organized lipids and marginally stable proteins: a coupling of weak interactions to optimize membrane signaling

Eukaryotic lipids in a bilayer are dominated by weak cooperative interactions. These interactions impart highly dynamic and pliable properties to the membrane. C2 domain-containing proteins in the membrane also interact weakly and cooperatively giving rise to a high degree of conformational plasticity. We propose that this feature of weak energetics and plasticity shared by lipids and C2 domain-containing proteins enhance a cell's ability to transduce information across the membrane. We explored this hypothesis using information theory to assess the information storage capacity of model and mast cell membranes, as well as differential scanning calorimetry, carboxyfluorescein release assays, and tryptophan fluorescence to assess protein and membrane stability. The distribution of lipids in mast cell membranes encoded 5.6-5.8bits of information. More information resided in the acyl chains than the head groups and in the inner leaflet of the plasma membrane than the outer leaflet. When the lipid composition and information content of model membranes were varied, the associated C2 domains underwent large changes in stability and denaturation profile. The C2 domain-containing proteins are therefore acutely sensitive to the composition and information content of their associated lipids. Together, these findings suggest that the maximum flow of signaling information through the membrane and into the cell is optimized by the cooperation of near-random distributions of membrane lipids and proteins. This article is part of a Special Issue entitled: Interfacially Active Peptides and Proteins. Guest Editors: William C. Wimley and Kalina Hristova.

Keywords: C2 domain Disorder Information theory Membrane domain Signaling.


Materials and Methods

Antibodies

The anti-RCP affinity-purified antibody has been described previously (Lindsay et al., 2002). The Rab11 and Rab11-FIP2 antibodies were raised in rabbits against Escherichia-coli-purified 6×His-Rab11 and 6×His-Rab11-FIP2(1-277). The resulting antisera were affinity purified and used at dilutions of 1:250 and 1:25, respectively. The anti-haemagglutinin (anti-HA) mouse monoclonal antibody was from Roche and used at a 1:1000 dilution. Mouse monoclonal anti-ZO1 antibody was purchased from BD Transduction Laboratories and used at a 1:150 dilution.

Plasmid constructs

Fusions between green fluorescent protein (GFP) and RCP and Rab11-FIP2 have been described previously (Lindsay et al., 2002 Lindsay and McCaffrey, 2002). GFP-Rip11 was generated by polymerase chain reaction (PCR) using pBluescriptIISK KIAA0857 (Kazusa DNA Research Institute) as template and the primers Rip11FWD (5′-CGGAATTCGCCCTGCGGGGCGCGGAG-3′) and Rip11REV (5′-CGGAATTCGCACAGAACCTGATGCCTCCAAG-3′). The product was digested and ligated into the EcoRI site of pEGFP-C1 (Clontech). GFP-RCP(1-199) was generated by ligating the ∼600 bp SalI fragment from pEGFP-C3 RCP into the SalI site of pEGFP-C3. GFP-RCP(200-649) was constructed by ligating the SalI-BamHI fragment from pEGFP-C3 RCP into the SalI-BamHI site of pEGFP-C1. GFP-Rab11-FIP2(1-284) was generated by PCR using Rab11-FIP2FWD (5′-CGGAATTCCCTGTCCGAGCAAGCCCAAAAG-3′) and Rab11-FIP2ΔC2REV (5′-CGGAATTCGAAACCAGCCACAGGATCAATTC-3′) and ligating into the EcoRI site of pEGFP-C1. GFP-Rab11-FIP2(277-512) was generated by ligating the HindIII-BamHI fragment from pEGFP-C1 Rab11-FIP2 into the HindIII-BamHI site of pEGFP-C2 (Clontech). GFP-Rip11(1-218) was generated by digesting pEGFP-C1 Rip11 with BamHI and religating the backbone. GFP-Rip11(219-653) was constructed by ligating the ∼2.2 kb BamHI fragment from pBSKII KIAA0857 into pEGFP-C1. RCP(1-199) was subcloned into pGEX2T (Amersham Biosciences). Rip11(1-218) was subcloned into the BamHI site of pGEX3X and Rab11-FIP2(1-284) was subcloned into the EcoRI site of pGEX3X (Amersham Biosciences). All constructs generated by PCR were confirmed by sequencing.

Expression and purification of GST fusion proteins

Glutathione-S-transferase (GST) fusion-protein expression was induced in E. coli XL-1 Blue cells with 0.1 mM isopropylthiogalactoside (IPTG) at 30°C for 4 hours. The bacterial cells were resuspended in 1× PBS and lysed with 1% Triton X-100 plus 5 mM dithiothreitol. The lysate was clarified by centrifugation at 12,000 g and the protein was bound to glutathione-agarose (Sigma), which was washed twice with PBS. The protein was eluted with 10 mM glutathione and subsequently dialysed overnight in PBS.

Protein-phospholipid assays

The protein-phospholipid overlay assays were carried out according to the manufacturers instructions. Briefly, PipArrays™ (Echelon Biosciences) were blocked in 3% bovine serum albumin (BSA) in Tris-buffered saline-0.1% Tween-20 (TBST) and then incubated overnight with 0.5 μg ml –1 GST fusion protein. The membranes were then washed extensively with 3% BSA in TBST and probed with anti-GST antibody (Amersham Biosciences). After washing, the membranes were incubated with horseradish peroxidase (HRP)-coupled anti-rabbit IgG and then visualized by autoradiography. Densitometry was performed using GeneTools analysis software (Syngene). The intensity of each phosphoinositide spot, at a concentration at which the signal was not saturated, was measured. The spot with the highest intensity was set at 100 [this was PtdIns(3,4,5)P3 in every case]. The intensity of the other phosphoinositide signals, at the same concentration, were normalized against PtdIns(3,4,5)P3. The results presented are the mean of three independent experiments. PipArrays with PA, phosphatidylcholine (PC), phosphatidylserine (PS) and phosphatidylethanolamine (PE) (Sigma) were made as described previously (Dowler et al., 2002).

Cell culture and immunofluorescence

HeLa and A431 cells were maintained in culture in Dulbecco's modified Eagle's medium (DMEM) (BioWhittaker) supplemented with 10% foetal bovine serum, 100 units ml –1 penicillin, 100 μg ml –1 streptomycin and 2 mM glutamine. Transfections were carried out using Effectene transfection reagent (Qiagen) according to the manufacturers instructions. 8 hours after transfection, the medium was replaced with fresh medium and, 16 hours later, the cells were fixed with 3% paraformaldehyde and mounted with Mowiol. For antibody labelling, the cells were permeabilized with 0.05% saponin supplemented with 0.2% BSA. Images were acquired on a confocal microscope (Zeiss LSM 510) using a PlanApo 63× 1.4 NA oil-immersion objective. Images were processed using Image Examiner software (Carl Zeiss) and imported into Adobe Illustrator.


Cell membranes and chemicals

Chemicals (foods, medicines, drugs of abuse, industrial chemicals) can enter the human body by various means including ingestion, inhalation, injection (intravenous, subcutaneous, intramuscular), skin application, use of a suppository and application to mucous membranes (eye, oral and nasal cavities). Except for injection directly into the bloodstream, the chemical must pass through a complex system of living cell membranes before it can enter the bloodstream.

For example, chemicals that enter the digestive tract must be absorbed by the cells lining the small intestine and then be transferred through the cell to the other side where the chemical can then be absorbed by the capillary cells into the bloodstream. Likewise, chemicals that are inhaled, as would occur from those released following the derailment, must pass through the alveolar cells to get to the capillary cells lying close to the alveoli to enter the bloodstream.

As chemicals pass into and out of cells, they must cross the cell membrane (Figure 1). It is the membrane that keeps all of the cell contents securely inside, but which allows some materials to pass in and out of the cell via several different mechanisms. The cell membrane consists mainly of phospholipid and protein in the form of a lipid bilayer. The two lipid layers face each other inside the membrane, and the more water soluble parts of the phospholipid molecule (phosphate groups) face the aqueous media inside the cell (cytoplasm) as well as outside the cell (intercellular fluid). The structural relationship of the proteins and phospholipids in the membrane was determined by two scientists, S.J. Singer and G. Nicolson, and is termed a "fluid mosaic model."

Figure 1: The Cell Membrane. Chemicals must pass through the membrane to enter or exit cells.

Chemicals can enter the body through our eyes.


Materials and methods

Mice were housed in constant temperature at 23 °C ± 1 °C with 40% humidity under a 12 h light–dark cycle. Eight-week-old female B6D2F1 [BDF1, cross between C57BL/6 J (B6) female x DBA/2 J (D2) male] mice were purchased from Orient Bio (Gyunggi-do, Korea). Mice between 10 and 14-weeks-old were used in the “young” group, whereas mice older than 45 weeks were used in the “old” group. All experiments were conducted in accordance with the policies of the Konkuk University Institutional Animal Care and Use Committee (Approval number KU17067). Mice were sacrificed under anesthesia, and all efforts were made to minimize suffering of the mice.

Oocyte collection

Mice received 10 IU of pregnant mare’s serum gonadotropin (PMSG, Sigma-Aldrich, St. Louis, USA) intraperitoneally, and 10 IU of human chorionic gonadotropin (hCG, Sigma-Aldrich) 48 h later (7–8 pm). Ovulated cumulus-oocyte complex (COCs) were retrieved from the oviduct 13 h post-hCG (8–9 am). 20 IU of PMSG and hCG injections were used for better induction in older mice, as in previously conducted research [10, 23]. To remove the cumulus cells, the COCs were treated with hyaluronidase (300 mg/ml, H4272, Sigma Aldrich) in Quinn’s Advantage Medium with HEPES (SAGE Media, ART-1023, Trumbull, CT, USA) for 2 min at room temperature. Denuded metaphase II (MII) oocytes were washed and collected in Quinn’s Advantage Medium with HEPES containing 20% fetal bovine serum (FBS, Gibco Grand Island, NY, USA).

Vitrification and warming

Vitrification was performed as previously described by Cha et al. [24]. Ethylene glycol (EG, 102466, Sigma-Aldrich) and dimethyl sulfoxide (DMSO, D2650, Sigma-Aldrich) were used as cryoprotectants in the vitrification solution. Oocytes were equilibrated in PBS based media containing 7.5% EG, 7.5% DMSO, and 20% FBS for 2.5 min, and then transferred to media containing 15% EG, 15% DMSO, and 0.5 M sucrose (Fisher Scientific, Fair Lawn, USA) for 20 s. Equilibrated oocytes (20 to 25) were loaded onto a copper grid (Ted Pella Inc., Redding, USA) and dipped directly into liquid nitrogen (LN2). Vitrified oocytes were stored in a LN2 tank for 2–4 weeks. For the warming procedure, the grid was taken out from the LN2 tank and serially incubated in 20% FBS containing PBS with descending concentrations of sucrose (0.5, 0.25, 0.125, 0 M) for 2.5 min each. The vitrified-warmed oocytes were washed in Quinn’s Advantage medium containing HEPES and 20% FBS. Washed oocytes were cultured in M16 media (M7292, Sigma-Aldrich) at 37 °C, in 5% CO2 for 1 or 3 h. For Necrostatin-1 (Nec1, N9037, Sigma-Aldrich) supplementation, 1 μM of Nec1 [25] was added to the final vitrification solution (15% EG, 15% DMSO, and 0.5 M sucrose). Oocytes without any marked morphological deformation and discoloration under an inverted microscope were considered as survived ones and used for further analysis.

RNA extraction and quantitative real-time polymerase chain reaction analysis

MII oocytes obtained from multiple mice were pooled and randomly grouped in 20 for RNA extraction. mRNA was extracted from 20 oocytes in all experimental groups, using Dynabeads™ mRNA DIRECT™ Purification Kit (Life Technologies, Forster City, CA, USA) according to the manufacturer’s protocol. The mRNA was kept in − 80 °C before use. First strand cDNA was synthesized from total mRNA sample using a Superscript TM III Reverse Transcriptase (Invitrogen, 18,080–044, Carlsbad, CA, USA), RNaseOUT™ Recombinant Ribonuclease Inhibitor (Invitrogen, 10,777–019), Oligo (dT)20, and random hexamer primers (Roche, Basel, Switzerland). Real-time quantitative PCR (qPCR) was performed using 2 μl of oocyte cDNA (equivalent to one oocyte per reaction) and Applied Biosystems™ Power Up™ SYBR™ Green Master Mix (Invitrogen, A25742, Carlsbad, CA, USA) in a final volume of 20 μl on the ABI 7500 real-time PCR system. The PCR conditions were as follows: hold for 10 min at 95 °C, followed by each cycle consisting of denaturation at 95 °C for 15 s, annealing and elongation at 58 °C for 1 min each. The relative gene expression was normalized with H2afz mRNA expression and relative quantification was performed using the ddCt method [26, 27]. PCR was performed by using Econo Taq PLUS GREEN 2X Master Mix (Lucigen, Middleton, WI, USA). Three biological replicates were used per experimental group and all reactions were run in duplicates. Primers used are shown in Table 1.

Cell culture and necroptosis induction

L929 fibroblast cell line derived from mouse adipose tissue was obtained from Korean Cell Line Bank (Seoul, Korea). L929 cells were cultured in RPMI1640 media supplemented with L-glutamine (300 mg/L), 25 mM HEPES, 25 mM NaHCO3, and 10% FBS. To induce necroptosis, cells were treated with a mixture of 30 ng/mL TNFα (PeproTech, Rocky Hill, NJ), 10 μM LCL-161 (R&D Systems, Minneapolis, MN, USA), and 20 μM Z-VAD-FMK (R&D Systems) for 40 min [28]. Cells were fixed and subjected to immunofluorescence staining with anti-pMLKL and anti-pRIPK1 antibodies to establish specificity of these antibodies.

Immunofluorescence staining

Oocytes were fixed with 4% paraformaldehyde containing 0.05% polyvinyl alcohol in phosphate buffered saline (PFA-PVA) for 10 min. Fixed oocytes were washed three times with phosphate-buffered saline containing 0.05% polyvinyl alcohol (PBS-PVA) for 10 min each. For permeabilization, oocytes were transferred to a solution containing 0.25% Triton X-100 and incubated for 10 min. To prevent nonspecific binding, oocytes were blocked with 2% BSA in PBS for 1 h, followed by incubation with primary antibody at 4 °C overnight. The primary antibodies used were anti-pMLKL (1:100, ab196436, Abcam) [21], anti-pericentrin (1:500, 611,814, BD Bioscience, San Jose, CA, USA) [29], and anti-pRIPK1 (1:150, 31,122, Cell Signaling Technology, Danvers, MA, USA). Following incubation with primary antibodies, oocytes were washed three times with PVA-PBS for 10 min. The oocytes were then incubated with Alexa Fluor 488 or Alexa Fluor 568 conjugated secondary antibody (1:250, Invitrogen) for 1 h at room temperature. DNA was stained with TOPRO-3-iodide (1:250, Invitrogen). The oocytes were mounted on glass slides with Vectashield mounting medium (Vector Laboratories, Peterborough, UK) and observed under the confocal microscope (Zeiss LSM710, Carl Zeiss AG, Oberkochen, Germany). Specificity of anti-pMLKL and anti-pRIPK1 antibodies was confirmed in necroptosis-induced L929 cells [21]. In all experiments, rabbit IgG was used as a negative control and it did not generate any specific signal.

Live imaging of oocytes by confocal microscopy

Oocytes were washed with M16 media three times and stained with CellMask™ Plasma Membrane Stain (2.5 μg/ml C10046, Life technologies), BODIPY 500/510 dodecanoic acid (10 μg/ml D-3823, Invitrogen), or ER Tracker™ Red dye (1 μg/ml E34250, Invitrogen) for 30 min. The oocytes were rinsed with fresh M16 media three times and transferred to a glass bottom confocal dish. Live images of oocytes were obtained directly with a confocal microscope (Zeiss LSM710).

Statistical analysis

Data analysis and graph preparation were done using GraphPad Prism 5 software. For statistical analysis, Student’s t-test or one-way analysis of variance (ANOVA) were conducted on the experimental groups. Tukey’s range test was then performed to identify whether a significant difference exists among the groups. Statistical significance was depicted as *: p < 0.05, **: p < 0.01 and ***: p < 0.001.


DISCUSSION

The distribution of aminophospholipids (PE and PS) in the membranes of Gram-positive bacteria was known to be asymmetric from previous experiments using TNBS probe. PE distribution was 67%/33% and 80%/20% favoring the cytoplasmic leaflet of Bacillus megaterium (12, 15) and eukaryotic cells (22), respectively. At the same time, the PE in Bacillus amyloliquefaciens was found to be located predominantly in the outer leaflet of cytoplasmic membrane (33). Although still controversial, these studies indicate that PE is asymmetrically distributed in bacterial membranes and raised the question of its transmembrane distribution in Gram-negative bacteria, which have a more complex envelope system of IMs and OMs. No similar investigations have been reported for Gram-negative bacteria. Thus, we exploited permeant and nonpermeant amino group–specific probes to determine the aminophospholipid head group distribution, followed by assessment of their acyl group asymmetries in the E. coli IM. Characterization of labeled lipids with independent radiolabeled, normal-phase LC/MS/MS, TLC elution–based, and TLC-less spectrophotometric assays allowed us to reveal asymmetrical, dynamic, and cell shape–dependent PE distribution in the IM of Gram-negative bacteria.

Our work represents a technical and conceptual breakthrough in the following areas:

1) Quantitative analysis of PE sidedness led to the first unambiguous demonstration of PE asymmetry in IM of Gram-negative bacteria such as E. coli (Figs. 2 and 3) and Y. pseudotuberculosis (fig. S4), which has long remained unknown.

2) Asymmetry of PE is a remarkable dynamic. PE appears first on the periplasmic side of the IM of E. coli cells, followed by dynamic and disproportional distribution to the cytoplasmic leaflet (Fig. 5).

3) Rod-shaped and filamentous E. coli cells display an opposite distribution of PE in their IMs (Figs. 3 and 4, E to H). This very intriguing finding suggests that PE distribution facilitates or results from changes in bacterial shape. The rates of de novo fatty acid and CL biosynthesis can dictate bacterial shape, as was demonstrated recently (34, 35). The rate of CL biosynthesis and cells shape of logarithmically grown E. coli cells are interrelated quantitatively (35). Gradual changes in distribution of PE/CL amounts between the IM leaflets during de novo PE biosynthesis in E. coli cells initially lacking PE coincides with progressive reduction of cell size (Fig. 5A). PE is progressively accumulated, but CL is dissipated from the cytoplasmic leaflet of the IM. These transmembrane redistributions of CL and PE correlate with gradual changes in lipid packing order predominantly in the luminal leaflet of ISOv (corresponding to periplasmic leaflet of IM) (Fig. 6, C and D), demonstrating the possible coinvolvement of all these events in bacterial size control.

4) Lipid asymmetry is most likely metabolically controlled, regulated, and maintained in the IM of Gram-negative bacteria. The transmembrane distribution of PE and PS presumably reflects the relative rates of synthesis, translocation, and utilization of these aminophospholipids and their metabolic precursors. The asymmetric distribution of PE in IM can result from the balance between its translocation to the cytoplasmic leaflet, biosynthetic buildup on this leaflet, and loss from the periplasmic leaflet due to biosynthetical utilization and flow to the OM (illustrated in Fig. 6E). This balance is likely to be different in rod-shaped and filamentous cells because of different relative areas of IM and OM and different dynamics of membrane growth. This may explain the opposite asymmetric distribution of PE in the IM of rod-shaped and filamentous cells (Fig. 4 and fig. S7, D and E illustrated in Fig. 6F).

5) At any given time, more and more PE molecules are required at the outer leaflet of IM to generate mature triacylated lipoprotein precursor (Lpp) and provide material for exclusive building of the PE-enriched periplasmic leaflet of the OM (Fig. 6E). This is consistent with initial appearance of nascent PE in the outer IM leaflet in all cell types examined (Fig. 5). Although it may be advantageous to have the precursor of PS initially present in the IM periplasmic leaflet, a puzzling aspect of PS localization and accumulation in this leaflet in the psd conditionally lethal mutant (Fig. 4, A to D, and fig. S7, B and C) can be explained by its inability to serve as an acyl donor in the reaction with mature Lpp catalyzed by apolipoprotein N-acyltransferase (Fig. 6E). Nevertheless these results pose a provocative question as to the localization of the active site of PS decarboxylase and necessitate anterograde and retrograde movement of PS and PE, respectively.

6) No changes in acyl group asymmetry of PE, PG, or PS were observed in the IM of wild-type E. coli and its lipid mutants (fig. S6). Our observations suggest rather an existence of a link between PE distribution, CL content, and lipid order within the IM. Since PE may increase the bilayer rigidity in the range comparable with that of cholesterol (36), our results suggest that bacteria may maintain membrane leaflet composition and the lipid order and rigidity not only by regulating the saturation state of the newly synthetized phospholipids, as widely accepted, but also by a fine-tuning of PE and CL content of individual leaflets. Thus, compositional asymmetries and physical order can be intrinsically coupled in bacterial membranes (Fig. 6, A to D, and fig. S8, A to D illustrated in Fig. 6G).

Although membrane asymmetry is largely related to lipid distribution, a full understanding of the physiological significance and detailed molecular mechanisms by which membrane phospholipid asymmetry is generated and maintained is lacking. In this work, we report novel methodology for determination of aminophospholipid asymmetry, which is based on complementary labeling of their head groups by two vectorial chemical probes, TNBS and DFDNB. To determine both head group and acyl asymmetry, the labeled lipids could be further characterized to determine head group and acyl asymmetry of aminophospholipids using radiometric, mass spectrometric methods, and spectrophotometric approaches. These methods are suitable for investigation of transbilayer dynamics, steady-state sidedness, and physiological significance of aminolipid asymmetry in Gram-negative (PE, PS, and ornithine lipids) and Gram-positive (O-lysylphosphatidylglycerol) bacteria and in any single-membrane cellular organelle (exosomes and apoptotic bodies).

Our results reveal unexpected opposite asymmetry of PE in the cytoplasmic and periplasmic leaflets of E. coli IM of rod-shaped and filamentous cells. PE is dynamically redistributed within the IM either reflecting or facilitating the changes in cell shape. In eukaryotic cells, lipid asymmetry is maintained by the interplay of glycerophospholipid flippases and scramblases, which have not been discovered in bacterial cells. Thus, bacterial cells have to rely on different mechanisms for maintaining the membrane asymmetry. One of such mechanism is the dynamic balance between the incorporation of phospholipids into the cytoplasmic leaflet (due to their continuous synthesis) and their removal from the periplasmic leaflet (due to transport to the OM or usage in metabolic pathways involving periplasmic lipoproteins). This may create a steady-state concentration gradient of the lipids resulting in IM lipid asymmetry. Still unknown mechanisms of anterograde transport from IM to OM may accommodate the rates of de novo glycerophospholipid biosynthesis and dictate bacterial size, as was proposed recently (37). Thus, PE asymmetry in the IM might originate metabolically and play a central role in regulating the size and morphology of Gram-negative bacteria along with balanced synthesis of certain cell cycle proteins (20).

We also revealed that compositional asymmetries and physical order can be intrinsically coupled in the biogenic IM. The change in transmembrane distribution of two nonbilayer prone lipids, PE and CL, is tightly coupled to adjust the collective physical properties of the membrane. It is possible to speculate that this fundamental feature may be a main architectural principle for most, if not all, living biological membranes.

Obviously, further studies are necessary to fully elucidate the mechanisms and physiological consequences of our findings. Nevertheless, our results provide a conceptual basis and experimental tools for studying the transbilayer dynamics and regulatory function of various aminolipids (PE, PS, O-lysylphosphatidylglycerol, ornithine lipids, and some sphingolipids) in different organisms. These approaches offer the possibility of investigating the sidedness of aminolipids and establishment of their head and acyl group asymmetries in different single-membrane systems ranging from intact cell membranes of bacteria to erythrocytes, cell-derived organelles, and liposomes.


Calcium and Lipid selectivity

C2 domains are unique among membrane targeting domains in that they show wide range of lipid selectivity for the major components of cell membranes, including phosphatidylserine and phosphatidylcholine. This C2 domain is about 116 amino-acid residues and is located between the two copies of the C1 domain in Protein Kinase C (that bind phorbol esters and diacylglycerol) (see PDOC00379) and the protein kinase catalytic domain (see PDOC00100). Regions with significant homology Δ] to the C2-domain have been found in many proteins. The C2 domain is thought to be involved in calcium-dependent phospholipid binding Ε] and in membrane targeting processes such as subcellular localisation. Although most C2 domains interact with the membrane (phospholipids) in a Ca 2+ -dependent manner, some C2 domains can interact with the membrane without binding to Ca 2+ . Similarly, C2 domains have been evolved to have different specificities for lipids. Many C2 domains such as synaptotagmin C2A, bind to anionic phospholipids (PS or PIP2 containing phospholipids). However, other C2 domains such as cPLA2-α C2 domain bind to zwitterionic lipids (e.g. PC). This diversity and selectivity in Ca 2+ and lipid binding suggest that C2 domains are evolved to have different functions. Ζ]


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