NMR-based molecular ruler for determining the depth of intercalants within the lipid bilayer

Chemistry and Physics of Lipids 184 (2014) 105–118

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Chemistry and Physics of Lipids journal homepage: www.elsevier.com/locate/chemphyslip

NMR-based molecular ruler for determining the depth of intercalants within the lipid bilayer Part III: Studies on keto esters and acids Michal Afri, Carmit Alexenberg, Pinchas Aped, Efrat Bodner, Sarit Cohen, Michal Ejgenburg, Shlomi Eliyahu, Pessia Gilinsky-Sharon, Yifat Harel, Miriam E. Naqqash, Hani Porat, Ayala Ranz, Aryeh A. Frimer * The Department of Chemistry, Bar-Ilan University, Ramat Gan 5290002, Israel

A R T I C L E I N F O

A B S T R A C T

Article history: Received 9 March 2014 Received in revised form 10 July 2014 Accepted 21 July 2014 Available online 24 July 2014

The development of “molecular rulers” would allow one to quantitatively locate the penetration depth of intercalants within lipid bilayers. To this end, an attempt was made to correlate the 13C NMR chemical shift of polarizable “reporter” carbons (e.g., carbonyls) of intercalants within DMPC liposomal bilayers – with the polarity it experiences, and with its Angstrom distance from the interface. This requires families of molecules with two “reporter carbons” separated by a known distance, residing at various depths/polarities within the bilayer. For this purpose, two homologous series of dicarbonyl compounds, methyl n-oxooctadecanoates and the corresponding n-oxooctadecanoic acids (n = 4–16), were synthesized. To assist in assignment and detection several homologs in each system were prepared 13C-enriched in both carbonyls. Within each family, the number of carbons and functional groups remains the same, with the only difference being the location of the second ketone carbonyl along the fatty acid chain. Surprisingly, the head groups within each family are not anchored near the lipid–water interface, nor are they even all located at the same depth. Nevertheless, using an iterative best fit analysis of the data points enables one to obtain an exponential curve. The latter gives substantial insight into the correlation between polarity (measured in terms of the Reichardt polarity parameter, ET(30)) and penetration depth into the liposomal bilayer. Still missing from this curve are data points in the moderate polarity range. ã 2014 Elsevier Ireland Ltd. All rights reserved.

Keywords: Liposome 13 C NMR ET DMPC Oxooctadecanoates Molecular ruler

1. Introduction The biological membrane is the outer envelope of the living eukaryotic cell and protects the cell by modulating the crossing of chemicals into and out of the cell. Determining the location of molecules within biological membranes is important for a variety reasons. Inter alia, it allows one to understand the connection between the depth/orientation of a substrate within a membrane and its reactivity. Secondly, it is helpful in gaining insight into the structure and dynamics of various lipids and protein–lipid

* Corresponding author at: The Ethel and David Resnick Chair in Active Oxygen Chemistry, The Department of Chemistry, Bar-Ilan University, Ramat Gan 5290002, Israel. Tel.: +972 3 5318610; fax: +972 3 7384053. E-mail addresses: [email protected] (M. Afri), [email protected] (C. Alexenberg), [email protected] (P. Aped), [email protected] (E. Bodner) , [email protected] (S. Cohen), [email protected] (M. Ejgenburg), [email protected] (S. Eliyahu), [email protected] (P. Gilinsky-Sharon) , [email protected] (Y. Harel), [email protected] (M.E. Naqqash), [email protected] (H. Porat), [email protected] (A. Ranz), [email protected] (A.A. Frimer). http://dx.doi.org/10.1016/j.chemphyslip.2014.07.007 0009-3084/ ã 2014 Elsevier Ireland Ltd. All rights reserved.

interactions. There are number of spectroscopic techniques that have been used to study the intercalation of various molecules and peptides into biomembranes or membrane-models. These include solid-state (Hong, 2006) or solution (Prosser et al., 2007; Evanics et al., 2006) NMR, ATR-FTIR (Mourelatou et al., 2011), fluorescence quenching (Postupalenko et al., 2011; Herenyi et al., 2009; Caputo and London, 2003; Menger et al., 2002; Huster et al., 2001) and EPR spectroscopy (Altenbach et al., 1994; Fanucci and Cafiso, 2006; Frazier et al., 2002; Hubbell and Altenbach, 1994; Nielsen et al., 2005). Despite, or perhaps, because of its relative lack of complexity, the liposome is often used as a simple model for biological membranes (Gregoriadis, 1984; Papahadjopoulos, 1978). For decades, biophysicists have used for this purpose DMPC (dimyristoyl phosphatidylcholine, di-14:0-PC), a lipid with saturated myristoyl (14:0) chains at the sn-1 and sn-2 positions. Its popularity as a model membrane system can be attributed to the fact that it is stable, inexpensive, and easy to obtain. Because it has fully saturated acyl chains, DMPC bilayers are robust and do not require special preparatory environments. Over the years, DMPC bilayers have been well characterized using a variety of physical

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techniques (Ku9 cerka et al., 2011; Jämbeck and Lyubartsev, 2012; Aussenac et al., 2003), and as a result, they have been used to study problems ranging from drug–membrane interactions (Komljenovic et al., 2010), to the enhancement of cholesterol flipflop (Ku9 cerka et al., 2010), to the difference between bulk water and interfacial polarity and pH (Voinov et al., 2009), to examining the location, behavior and antioxidant properties of a-tocopherol in biomimetic membranes (Takahashi et al., 1989; Serbinova et al., 1991; Wassall et al., 1991; Afri et al., 2004a; Fukuzawa, 2008). Despite its broad use, there have been some authors who have questioned DMPC’s biological relevance due to its oxidative stability (Marquardt et al., 2014) and the low level of 14:0 fatty acid-containing phospholipids in biological systems (e.g., <1% in lipoproteins) (Ruíz-Gutiérrez et al., 1993). In light of the above wide application of DMPC liposomes, developing convenient and readily available techniques for determining the depth (i.e., the distance from the water–lipid interface) of substrates intercalated (henceforth “intercalants”) within the DMPC liposomal bilayer seemed of great value. In previous work, we have described an NMR method for this purpose which is based on two observations (Frimer et al., 1996; Afri et al., 1992, 2004a,b; Cohen et al., 2008a,b,c; Shachan-Tov et al., 2010). Firstly, there is a generally excellent linear positive correlation between solvent polarity and the 13C chemical shift (d) observed for a polarizable carbon (e.g., carbonyls). Perhaps the most famous polarity measure is the dielectric constant e (March, 1992) which expresses the polarizability and the ordering of the dipole of the solvent itself. This is in contradistinction to Reichardt’s ET(30) polarity parameter which is based on the interaction between the solvent and a polarizable solute (Reichardt, 1965, 1994; Reichardt and Welton, 2011). In particular, it measures the solvent dependency of the UV absorption (in kcal/mol at 25
C, 1 atm) of 2,6-diphenyl-4-(2,4,6-triphenyl-N-pyridine)-phenolate, dubbed Reichardt’s dye (Fig. 1). The more polar the solvent, the shorter the wavelength and the greater is its ET(30) value. Gamliel (2005) examined the 13C chemical shifts of the carbonyl of acetone, and correlated them with either the dielectric constant e or ET(30) polarity parameter over a range of 26 solvents. A substantially better correlation (R2 = 0.80) was obtained using the ET(30) polarity parameter than with the dielectric constant (R2 = 0.55), and hence the former was preferred for our future studies. In addition to the chemical shift- ET(30) solvent polarity correlation, the aforementioned NMR technique is based on a second important observation. A polarity gradient exists within the lipid membrane, ranging from that of water (ET(30) = 63.1 kcal/mol) at the water–lipid interface, down to hexane deep in the lipid slab (ET(30) = 31.0 kcal/mol). The extent of the ET(30)–d correlation is represented by the square of the correlation coefficient (R2), and experience has taught that it is generally quite high, R2 > 0.9 (Cohen et al., 2008a). This solvent polarity–chemical shift correlation can be readily attributed to the fact that carbonyls (and other polarizable moieties,

mutatis mutandis) have two resonance forms (Fig. 2), the non-polar double-bonded form and the charge-separated zwitterionic form. As the solvent polarity increases, the zwitterionic form – with lower electron density on the carbonyl carbon – makes a concommitantly greater contribution, and the NMR chemical shift of the carbonyl carbons increases, i.e., moves downfield. Due to the polarity gradient, the chemical shift values of a polar carbon within an intercalant will differ depending on its depth within the bilayer. Using the aforementioned correlation between the 13C chemical shifts and solvent polarity, one can prepare a “d/ET (30) correlation graph” for each of the polarizable “indicator” carbons in the molecule. When the molecule is then intercalated into the liposomal bilayer, the chemical shift measured reflects the micropolarity that carbon is experiencing. (Afri et al. (2002) have previously verified that with an intercalant concentration of 0.05 M and a intercalant:lipid molar ratio of 1:5, the intercalant itself has only a negligible affect on the polarity gradient. In addition, cryoTEM studies confirm that under these conditions the liposomes remain essentially unchanged (Afri et al., 2004a; Cohen et al., 2012).) Interpolating backwards from the correlation graph, converts the chemical shift value back into a polarity values, which allows us to approximate the extent of intercalant penetration into the bilayer. (The error in the ET(30) value is ca. 1.0 kcal/mol (Cohen et al., 2008a).) It should be emphasized that these penetration values are averages reflective of a dynamic residency of the probe molecule or intercalant. The timescale of molecular motion is faster than that of the NMR experiment; the NMR chemical shift data will reflect polarity due to electrostatic fields from the time-averaged structure, as well as from water penetration and perhaps partial exit of the surfactants. Despite this caveat, we believe that this 13C NMR chemical-shift/polarity correlation technique is a useful tool for approximating the location of substrates within lipid bilayers. Perhaps the major strength of the “NMR-technique” for locating intercalants in a bilayer is that it allows focusing on a specific atom, e.g., the carbon of a carbonyl (CQO) or nitonyl (CQN–R). Other techniques (like UV or fluorescence) require much lower intercalant concentrations, but focus on large chromophores; hence, positioning of the moiety can be tricky. The NMR concentration issue can be circumvented to some extent by 13C labeling. The NMR technique has allowed us to approximate the location of various substrates, such as coumarins (Afri et al., 2002), vitamin E (Afri et al., 2004a), dichlorofluorescein (Afri et al., 2004b), ubiquinone (Afri et al., 2004a) and curcumin (Afri et al., 2011), within DMPC liposomes. But these results are only qualitative in nature. It would be desirable to “translate” these qualitative ET(30) values to quantitative Angstrom depths from the interface. For the purpose of developing a convenient chemical ruler for DMPC liposomes, we turned to the synthesis (outlined in Fig. 3 and discussed below in Section 3) of two homologous series of dicarbonyl compounds on an 18-carbon fatty acid chain: methyl n-oxodecanoate esters 7, and the corresponding n-oxodecanoic acids 8. The numbering of the various carbons these dicarbonyl compounds is exemplified by 7-oxo analogs, ester 7d and acid 8d. In these bifunctional systems, one carbonyl is that of the acid or ester moiety, while the second is a ketone carbonyl moiety located

Double-bond O R C R'

Zwitterion O R C R'

Increasing solvent polarity Increasing chemical shift Fig.1. Reichardt’s dye.

Fig. 2. Resonance forms of the carbonyl.

M. Afri et al. / Chemistry and Physics of Lipids 184 (2014) 105–118

107

Fig. 3. General synthesis for keto esters 7 and keto acids 8. (*Position of possible 13C-labeling). (A) K*CN, DMSO, 75
C, overnight; (B) 40% KOH in EtOH, rt, overnight; (C) MeOH, H2SO4 (cat.), sonication overnight; (D) Ba(OH)28H2O, MeOH or EtOH, rt, overnight; (E) SOCl2, dry CH2Cl2, 30
C, 1h; (I) KOH 2.5N, EtOH, rt, overnight; (J) n = 4; MeOH, rt, 12h; n = 5: MeOH and CHCl3, reflux, 5h.

modularly along the chain at carbons 4–16. Based on literature precedent (Bronshtein et al., 2004), it was expected that the polar head groups of these bifunctional compounds would be anchored at or near the lipid–water interface. The keto-groups, on the other hand, are located at various locations along the lipophilic carbon chains and sense various polarity environments. Because of van der Waals force, these long chain compounds presumably lie parallel to the DMPC lipid chains, and the average Angstrom distance between the two carbonyls can be readily calculated. This location can then be correlated with the difference in ET(30) polarity of these same carbonyls. The data should permit the effective mapping of changes in polarity as one proceed further and further down the chain of the liposomal bilayer. The information obtained should enable one to construct a molecular ruler, complementing previously reported preliminary work in this field (Cohen et al.,

2008a). We chose an 18-carbon chain ruler so that we could scan the 14-carbon myristic chain fully and still have a portion of the carbon beyond the second DMPC carbonyl, to keep the tail aligned with the lipid chains. The general synthesis of dicarbonyls 7 and 8, outlined in Fig. 3, allows the preparation of derivatives a, e and h with the carbonyl moieties doubly enriched with 13C. This proved to be a great boon in detection and peak assignment. 2. Materials and methods 2.1. General Nuclear magnetic resonance (NMR) spectra were recorded on a variety of Bruker Fourier transform spectrometers: DRX 200 MHz,

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M. Afri et al. / Chemistry and Physics of Lipids 184 (2014) 105–118

DRX 300 MHz, DMX 600 MHz or Avance III 700 MHz instruments. For 1D NMR spectra, a QNP probe was used. All 2D experiments (COSY, HMQC, HMBC, and NOESY) were run and processed by Bruker software. NMR spectra were generally run at 25  1
C except in the case of DMPC liposomes, which were run at physiological temperature 37  1
C, above the phase transition temperature (Tm) of DMPC which is 23.6
C (Sassaroli et al., 1990). 1 H and 13C NMR chemical shifts are expressed in d (ppm) relative to TMS (0 ppm). Generally the sample was calibrated by internal TMS, though sometimes relative to the solvent. In the case of aqueous liposome solutions, the spectrum was calibrated according to the trimethylammonium peak at 54.6 ppm. In the 13C NMR spectra of the long chain derivatives, recorded as ppm relative to TMS, there are overlap peaks; therefore, the number of the chemical shifts does not always match the number of carbons. In addition, in the hydrogen and carbon assignments below, the various carbons were generally numbered according to the IUPAC nomenclature rules. Mass spectra (MS) and high resolution mass spectra (HRMS) were collected in a Waters (UK) AutoSpec Premier instrument in DCI/CH4 or CI/CH4 mode or by Q TOF in ESI mode. MALDI Mass spectra were obtained on with a Bruker (Germany) Autoflex III TOF/TOF machine. Infrared Fourier transform spectra (FTIR) were obtained using a Bruker Vector 22 instrument, with the data processed by Opus 5.0 (Bruker) software. The spectra were analyzed with IR MentorPro 6.5 (Bio-Rad) software. Abbreviations used for reporting % transmittance were: s = strong (100–75%), m = medium (75–45%), w = weak (45–25%), v = very and b = broad. Melting point was run on a Uni-Melt Thomas Hoover-capillary melting point apparatus manufactured by Thomas (Philadelphia PA, USA). Sonications were carried out either in a bath sonicator (EW-08890-06Cole-Parmer1 ultrasonic cleaner, 1/2 gallon, 230 VAC), or using a probe sonicator (Vibra-CellTM VCX130 Sonics and Materials Inc., ultrasonic processor with 20 kHz output frequency and a titanium alloy Ti-6Al-4V probe). The former was used for esterification reactions, while the later was used for liposome formation. Other standard equipment utilized included a vortex >(Winn Vortex Genie). The correlation coefficient (R2) for the chemical shift-solvent polarity correlation graphs were calculated by excel. Ab initio calculations were performed with Gaussian 03 performed at the B3LYP level using 6–31G* basis set Gaussian, Inc., Pittsburgh PA, 2003. Molecular modeling calculations were carried out with PCMODEL version 7.50.00, Serena Software, Bloomington, Indiana, USA – which uses the MMX force field. 2.2. Chemicals Acetyl chloride, 1-bromohexane, 1-bromodecane, EDTA, HEPES, KCN, Mg, dimyristoyl phosphatidylcholine (DMPC), methyl iodide, octyl bromide, octyl magnesium bromide, adipic acid, palmitic acid, sebacic acid, suberic acid, succinic acid, thionyl chloride, trifluoroacetic anhydride, anhydrous chloroform, anhydrous DMSO and the deuterated solvents were obtained from Sigma–Aldrich. Bromotetradecane, dibromodecane, dodecyl bromide and Tris–HCl were purchased from ACROS. Dibromohexane and adipoyl chloride methyl ester were acquired from Alfa Aesar. Barium hydroxide octahydrate (Fluka), sebacic acid monomethyl ester (ABCR Chemical Co.), cadmium chloride (Strem Chemicals), sodium azide (Fisher Scientific) and extra dry diethyl ether (Bio Lab Co.), were also commercially available. Regarding the 13C-labeled reagents, acetyl chloride-1-13C, K13CN and succinic anhydride-1 ,4-13C2 were obtained from Sigma–Aldrich, octanoic acid-1-13C and palmitic acid-1-13C were acquired from ISOTEC, and succinic acid1,4-13C2 was purchased from Cambridge Isotope Laboratories, Inc. Phosphate buffer saline solution pH 7.4 (PBS-N3) was prepared using doubly distilled water (dd H2O – purified via Millipore

Milli-Q columns) containing 1.7 mM NaH2PO4, 8.1 mM Na2HPO4, 2.7 mM KCl, 137 mM NaCl and 1 wt% sodium azide. 2.3. Sonic intercalation of compounds into DMPC liposomal solutions All glassware was first rinsed with conc. HCl to remove all traces of detergents and then with doubly distilled water. In a typical liposome preparation, the compounds to be intercalated (henceforth dubbed “intercalants”) and DMPC in a molar ratio of 1:5 were dissolved in chloroform in a vial. The solvent was then evaporated with a gentle steam of N2 while rotating in the palm of the hand, leaving a uniformly thin layer of lipid on the walls of the vial. The vial was finally set in a cotton-packed RB flask and the solvent was removed by rotary evaporation, leaving a uniformly thin film layer of lipid on the walls of the vial. The vial was then charged with 0.1 M phosphate buffer with 10% D2O (pH 7.8) yielding a cloudy solution in the desired molar concentration which was generally 0.05 M in intercalant. The lipid solution was vortexed for 10 min to obtain multilamellar liposomes. The liposomal solution was then sonicated until a clear solution of essentially unilamellar liposome was obtained. Previous cryo-TEM work (Afri et al., 2004a) confirms that these conditions produce primarily unilamellar liposomes. NMR of the liposomal solution indicates that the DMPC and intercalants remained essentially inert throughout the sonication, undergoing neither oxidation or hydrolysis. To verify that the various intercalants lie within the lipid bilayer and not in the aqueous phase, the liposomes were centrifuged down (25,000  g for 15 min) to a lipid pellet. The supernatant liquid was decanted and replaced by buffer, and the pellet was redispersed by vortexing the sample. NMR spectra of the starting and final liposomal and supernatant solutions revealed that the substrate indeed resides within the lipid bilayer exclusively. 2.4. General synthesis of keto esters 7 and keto acids 8 Many of the compounds described below are commercially available or have been previously reported. Nevertheless, we describe the synthesis and report spectral data in those cases where the 13C-enriched analogs were prepared in order to achieve sharp clear NMR spectra. Moreover, for many of these compounds, the complete spectral data (including splitting constants and assignments – which are far from trivial) are missing in the literature, or were previously measured with low resolution instruments. The NMR spectral data and assignments appear in Tables 1–5 while the complete spectral data including integration and splitting constants appear in the Supplementary information. 2.4.1. General preparation of alkanedinitriles 2 1,n-Dibromoalkane 1 were reacted with KCN (or K13CN) in dry DMSO following the procedure of Holmes and Lightner (1996); yielding the corresponding 1,n + 2-alkanedinitriles in generally excellent yields. The unlabeled analogs are commercially available (Sigma–Aldrich). 2.4.1.1. Octanedinitrile-1,8–13C2 (13C-2e). 100% yield. MS (DCICH4) m/z: 139.1234 (MH+, 4.49%), 111.1143 (M+–HCN, 27.44%), 97.0996 (M+  HCNCH2, 100%), 83.0871 + (M –HCN C2H4, 33.61%), 69.0887 (M+–HCNC3H6, 66.28%). HRMS (DCI CH4) m/z: calcd. (12C613C2H13N2, MH+) 139.1235, obsd. 139.1234. 2.4.1.2. Dodecanedinitrile-1,12-13C2 (13C-2h). 100% yield. MS (DCICH4) m/z: 195.178 (MH+, 100%), 167.160 (MH+ CH2N, 13.39%). HRMS (DCI CH4) m/z: calcd. (12C1013C2H21N2, MH+) 195.1772, obsd. 195.1777.

M. Afri et al. / Chemistry and Physics of Lipids 184 (2014) 105–118

109

Table 1 1 H NMR data for compounds 2–6.

H-2 H-3 H-4 H-5 H-6 H-7 H-8 H-9 H-10 H-11 H-12 H-13 H-14 H-15 OH OCH a b

2ea

2ha

3ea

3ha

4ea

4ha

5aa

5ha

5i

5j

5k

6c

6d

6ea

6f

6ha

6i

2.38 1.88b 1.51 1.51 1.69b 2.38

2.35 1.64 1.44 1.31 1.31 1.31 1.31 1.44 1.64 2.35

2.29 1.60 1.37 1.37 1.60 2.29

2.28 1.56 1.32 1.32 1.32 1.32 1.32 1.32 1.56 2.28

2.31 1.63 1.34 1.34 1.63 2.31

2.30 1.62 1.28 1.28 1.28 1.28 1.28 1.28 1.62 2.30

2.32 1.63 1.35 1.35 1.63 2.35

2.30 1.62 1.28 1.28 1.28 1.28 1.28 1.28 1.62 2.34

2.23 1.59 1.19 1.19 1.19 1.19 1.19 1.19 1.19 1.59 2.26 11.12

2.31 1.60 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.60 2.31

2.35 1.73 1.73 2.92

2.27 1.64 1.37 1.64 2.85

2.22 1.54 1.27 1.27 1.63 2.81

2.29 1.60 1.31 1.31 1.31 1.69 2.86

2.30 1.61 1.28 1.28 1.28 1.28 1.28 1.28 1.70 2.86

2.27 1.58 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.67 2.85

2.83

2.85 3.67

3.67

3.67

3.67

3.59

3.65

2.29 1.59 1.22 1.22 1.22 1.22 1.22 1.22 1.22 1.22 1.22 1.22 1.59 2.29 10.81 3.63

3.68

3.56

3.57

3.56

3.66

3.63

13

These compounds were also prepared with C-enriched nitriles or carbonyls. The assignments for these absorptions are indefinite and may be interchangeable with each other.

2.4.2. General preparation of diacids 3 The alkanedinitriles 2 were hydrolyzed to the corresponding diacids using 40% KOH in ethanol, according to the procedure Holmes and Lightner (1996). The unlabeled analogs 3 are commercially available (Sigma–Aldrich). 2.4.2.1. Octanedioic acid-1,8-13C2 (13C-3e). 77% yield; mp: 143
C. MS (DCI CH4) m/z: 176.093 (M, 6.18%), 159.094 (MH+ H2O, 100%), 130.093 (M H2O CO, 20.62%). HRMS (DCI CH4) m/z: calcd. (12C613C2H14O4, M) 176.0959, obsd. 176.0926. 2.4.2.2. Dodecanedioic acid-1,12-13C2 (13C-3h). 99% yield; mp: 127–129
C. MS (DCICH4) m/z: 233.161 (MH+, 37.36%), 215.158 (MH+ H2O, 100%). HRMS (DCI CH4) m/z: calcd. (12C1013C2H23O4, + MH ) 233.1663, obsd. 233.1613. 2.4.3. General preparation of dimethylesters 4 Diesters 4 were prepared according to the procedure described by Khurana et al. (1990), by sonicating the appropriate dicarboxylic acid with methanol and catalytic sulfuric acid. The unlabeled analogs 4 are commercially available (Sigma–Aldrich).

2.4.3.1. Dimethyl octanedioate-1,8-13C2 (13C-4e). 81% J = 57.5 Hz, C2 and C7), 28.82 (d, J = 3.5 Hz, C4 and C5), (d, J = 3.0 Hz, C3 and C6). MS (DCI CH4) m/z: 205.135 49.56%), 173.106 (MH+ CH3OH, 100%), 130.086 +  CO2CH3 CH3 + H+, 19.72%). HRMS (DCI CH4) m/z: (12C813C2H19O4, MH+) 205.1350, obsd. 205.1346.

yield. 24.81 (MH+, (MH calcd.

2.4.3.2. Dimethyl dodecanedioate-1,12-13C2 (13C-4h). 45% yield, mp: 30–32
C. MS (DCI CH4) m/z: 275.203 (MH+ + CH5, 6.67%), 261.197 (MH+, 11.87%), 229.172 (MH+ CH3OH, 100%). HRMS (DCI CH4) m/z: calcd. (12C1213C2H27O4, MH+) 261.1976, obsd. 261.1965. FTIR (KBr): 2919, 2851, 1730 (s, CQO) cm1. 2.4.4. General procedure for preparation of monoester monoacid 5 Monomethyl succinate (5a) was prepared from succinic anhydride (13a, 0.5 g) (or the commercially available (Sigma–Aldrich) 1,4-13C2 labeled analog) and methanol (11 mL) according to the procedure of Lodyato et al. (2004). Monomethyl glutarate (5b) was prepared from glutaric anhydride (13b) and methanol according to the procedure described by Oda et al. (1995). The remaining monoesters were prepared from the corresponding diesters 4 via monohydrolysis with Ba(OH)28H2O

Table 2 1 H NMR data for keto esters 7 and keto acids 8.

H-2 H-3 H-4 H-5 H-6 H-7 H-8 H-9 H-10 H-11 H-12 H-13 H-14 H-15 H-16 H-17 H-18 OH OCH a b c

7aa

7c

7ea

7ha

7i

7k

8aa

8b

8c

8d

8ea

8f

8g

8ha

8i

8j

8k

2.72b 2.59b –c 2.44 1.56 1.25 1.25 1.25 1.25 1.25 1.25 1.25 1.25 1.25 1.25 1.25 0.88

2.38 1.62 1.62 2.38 –c 2.38 1.62 1.26 1.26 1.26 1.26 1.26 1.26 1.26 1.26 1.26 0.88

2.22 1.54 1.19 1.19 1.54 2.25 –c 2.25 1.54 1.19 1.19 1.19 1.19 1.19 1.19 1.19 0.91

2.30 1.61 1.27 1.27 1.27 1.27 1.27 1.27 1.56 2.38 –c 2.38 1.56 1.27 1.27 1.27 0.88

2.30 1.58 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.58 2.38 –c 2.38 1.58 1.58 1.24 0.89

2.39 1.59 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.59 2.39 –c 2.39 1.04

2.71b 2.64b –c 2.44 1.53 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.24 0.86

2.42 1.90 2.39 –c 2.39 1.55 1.25 1.25 1.25 1.25 1.25 1.25 1.25 1.25 1.25 1.25 0.88

2.41 1.62 1.62 2.41 –c 2.41 1.62 1.25 1.25 1.25 1.25 1.25 1.25 1.25 1.25 1.25 0.88

2.37 1.58 1.24 1.58 2.37 –c 2.37 1.58 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.24 0.86 9.67

2.28 1.51 1.19 1.19 1.51 2.32b –c 2.31a 1.51 1.91 1.91 1.91 1.91 1.91 1.91 1.91 0.81

2.33 1.52 1.26 1.26 1.26 1.52 2.33 –c 2.33 1.52 1.26 1.26 1.26 1.26 1.26 1.26 0.88

2.34 1.59 1.27 1.27 1.27 1.27 1.59 2.38 –c 2.38 1.59 1.27 1.27 1.27 1.27 1.27 0.88

2.28 1.54 1.30 1.30 1.30 1.30 1.30 1.30 1.54 2.28 –c 2.28 1.54 1.30 1.30 1.30 0.87

2.26 1.51 1.25 1.25 1.25 1.25 1.25 1.25 1.25 1.51 2.31 –c 2.31 1.51 1.18 1.18 0.81 10.64

2.36 1.55 1.25 1.25 1.25 1.25 1.25 1.25 1.25 1.25 1.55 2.36 –c 2.36 1.55 1.25 0.89

2.38 1.59 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.24 1.59 2.38 –c 2.38 1.04

3.68

3.67

3.6

3.66

3.66

3.66

13

These compounds were also prepared with C-enriched carbonyls. The assignments for these absorptions are indefinite and may be interchangeable with each other. Carbonyl position.

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M. Afri et al. / Chemistry and Physics of Lipids 184 (2014) 105–118

Table 3 13 C NMR data for compounds 2–5.

C-1 C-2 C-3 C-4 C-5 C-6 C-7 C-8 C-9 C-10 C-11 C-12 C-13 C-14 C-15 C-16 OCH a b c d e f

2ea

2ha

3ea

3ha

4ea

4ha

5aa

5ha

5i

5j

5k

119.57 17.08 25.08 27.87 27.87 25.08 17.08 119.57

119.86 17.07 25.31 28.58 29.11 28.66 28.66 29.11 28.58 25.31 17.07 119.86

174.63 34.11 25.50 29.52 29.52 25.50 34.11 174.63

174.62 34.16 25.66 29.84 30.18c 30.04c

174.22 34.04 24.81 28.82 28.82 24.81 34.04 174.22

174.39 34.17 25.03 29.21 29.44c 29.30c

174.03 34.10 24.85 28.86 28.86 24.85 34.00 179.62

174.51 34.17 25.01 29.19 29.41b 29.27 29.27 29.40b 29.11 24.75 34.14 180.11

174.5 34.13 24.97b 29.51d 29.42d 29.26d 29.16d 25.08d

174.59 34.21 25.06b 29.64e 29.52e 29.34e 29.25e 25.16e

174.55 34.18 25.02b 29.66f 29.51f 29.32f 29.22f 25.14f

29.84 25.66 34.16 174.62

29.21 25.03 34.17 174.39

51.53

51.50

51.38

51.53

24.71b 34.13 180.21

24.77b 34.21 180.4

51.50

51.60

24.75b 34.18 180.13 51.54

13

These compounds were also prepared with C-enriched nitriles or carbonyls. The assignments for these absorptions are indefinite and may be interchangeable with each other. These values were observed for C-5–C-8. Five values were observed for C-4–C-10. Five values were observed for C-4–C-11. Five values were observed for C-4–C-13.

in MeOH or EtOH at room temperature overnight, following Tanaka (1959) and Rao et al. (1987). The unlabeled analogs 5a–h are commercially available (Sigma–Aldrich). 2.4.4.1. Monomethyl butanedioate-1,4-13C2 (13C-5a). 98% yield, mp: 55–59
C. HRMS (DCI CH4) m/z: calcd. (12C313C2H9O4, MH+) 135.0568, obsd. 135.0572. 2.4.4.2. Monomethyl octanedioate-1,8-13C2 (13C-5e). 26% yield, mp: 17–19
C. MS (DCI ICH4) m/z: 173.1043 (M+ OH, 9.96%), 159.0924 (M+OCH3, 35.35%), 140.0761 (MH+CH3OH H2O, 87.66%). HRMS (DCI CH4) m/z: calcd. (12C713C2H15O3, M+ OH) 173.1088, obsd. 173.1043.

2.4.4.3. Monomethyl dodecanedioate-1,12-13C2 (13C-5h). 51% yield, mp 50–52
C. MS (DCI CH4) m/z: 247.185 (MH+, 9.52%), 229.172 (MH+ H2O, 100%), 215.159 (MH+CH3OH, 30.33%). HRMS (DCICH4) m/z: calcd. (12C1113C2H25O4, MH+) 247.1820, obsd. 247.1851. 2.4.4.4. Monomethyl tridecanedioate (5i).

72% yield, mp: 50–52
C.

2.4.4.5. Monomethyl tetradecanedioate (5j). 75% yield. MS (CI, CH4) m/z 273.203 (MH+, 5.67%), 255.193 (MH+ H2O, 100%), 241.178 (MH + MeOH, 47.66%); anal. HRMS (CI): calcd. (C15H29O4, MH+) 273.2066 obsd. 273.2032. FTIR (KBr) 3000 (vbr, s, OH), 2912, 2845, 1737 and 1710 (s, CQO) cm1.

Table 4 13 C NMR data for compounds 6–7. 6c

6d

6ea

6f

6ha

6i

7aa

7c

7ea

7ha

7i

7k

C-1 C-2 C-3 C-4

173.53 33.41 23.66 24.48

173.73 33.52 24.67b 27.81

173.81 33.69 24.44 28.40b

174.26 34.07 25.06b 28.81c

173.89 33.87 24.49 23.22

174.32 34.1 24.91 29.47b

174.36 34.15 25.01 29.32b

46.71

24.30b

27.90b

28.89

42.98

42.90b

29.41b

29.78c

173.4 34.26 25.09 29.64j 29.54j 29.40j 29.28j

174.53 35.99b 25.10c 29.74k 29.57k 29.42k 29.28k

C-6 C-7 C-8 C-9 C-10 C-11 C-12 C-13 C-14 C-15 C-16 C-17 C-18 OCH3

173.45 51.69

46.79 173.59

24.71 46.83 170.5

28.31b 24.87c 47.15 173.91

174.35 34.14 25.11b 29.47c 29.41c 29.33c 29.27c 29.17c 29.09c 28.46c

173.46 22.87 37.16 209.34

C-5

174.35 34.09 25.06b 29.30c 29.25c 29.21c 29.12c 29.05c 28.41c

23.98 29.36 29.55 29.85g

210.67 42.27b 23.89 29.64h 29.48h 29.43h 29.36h 29.28h

23.67 42.87 211.69 42.84c 23.67 42.87c 29.84i 29.80i 29.67i

29.45d 29.29d 29.74c 29.20b 23.94e 42.87f 211.71 42.84f 23.93e 29.02b 31.69 22.57 14.08 51.46

24.03b 42.96 211.91 42.96 23.72b 31.59 26.6 14.06 51.53

24.11c 41.6 212.26 34.26b 7.99 51.58

24.94b 47.13 173.83

51.69

51.49

51.33

51.6

51.48

25.00b 47.17 173.9

51.46

29.65 32.08 22.85 14.25 51.93

31.93 22.69 14.13 51.54

32.03 22.82 14.25 51.61

b–f: The assignments for these absorptions are indefinite and may be interchangeable with each other in a given set for a given compound. a The starred compounds were also prepared with 13C-enriched nitriles or carbonyls. g This value corresponds to C-9–C-14. h C-9–C-15 corresponds to 5 values. i C-12–C-15 corresponds to three values. j C-4–C-10 show only 5 values. k C-14–C-14 show only 3 values.

M. Afri et al. / Chemistry and Physics of Lipids 184 (2014) 105–118

111

Table 5 13 C NMR data for keto acids 8.

C-1 C-2 C-3 C-4 C-5 C-6 C-7 C-8 C-9 C-10 C-11 C-12 C-13 C-14 C-15 C-16 C-17 C-18

8aa

8b

8c

8d

8ea

8f

8g

8ha

8i

8j

8k

178.22 27.86 36.91 209.33 42.90 23.95 29.79e 29.75e 29.60e 29.52e 29.49e 29.32e

177.21 32.71 18.63 42.94b 210.43 41.31b 23.87 29.69f 29.66f 29.62f 29.48f 29.42f 29.36f 29.26f

178.35 33.60 24.21 23.99c 24.92b 210.97 42.21b 23.88c 29.71g 29.66g 29.48g 29.42g 29.36g 29.26g

179.66 34.02 24.6 29.62b 29.61b 24.02c 43.00d 211.78 42.74d 23.71c 29.55i 29.40i 28.95b,i

178.80 33.78 24.59 29.42b,j

180.15 34.17 24.74 29.78o 29.57o 29.29o 29.11o

179.64 34.12 24.81 29.66p 29.54p 29.39p 29.35p 29.18p

24.00 42.96 212.06 42.96 24.00 29.09b 31.76 22.65 14.19

179.77 34.14 24.82b 29.73q 29.51q 29.55q 29.42q 29.37q 29.17q 29.03q

31.93 23.08 14.13

179.84 34.10 24.76 29.84l 29.52l 29.42l 29.31b,l 24.04c 42.99d 211.96 42.90d 23.95 29.18m 29.11m 29.05b,m 31.96 22.78 14.23

179.23 34.02 24.80 29.49n 29.38n 29.32n 29.15b,n

31.93 22.70 14.13

179.75 33.92 24.55 29.82b 24.02c 43.02d 211.54 42.54d 23.47c 29.73h 29.60h 29.54h 29.45h 29.39h 28.72b,h 32.03 22.81 14.23

23.95b 42.87c 212.1 42.84c 23.64b 31.15 22.54 13.91

32.06 22.83 14.26

32.02 22.80 14.24

23.89c 43.85d 211.67 42.70d 23.73c 29.26k 29.01k 28.84b,k 31.85 22.65 14.08

24.03 42.97b 212.04 42.67b 26.13 22.52 14.01

24.11b 42.61 212.42 36.00 8.00

b–d: The assignments for these absorptions are indefinite and may be interchangeable with each other in a given set for a given compound. a These compounds were also prepared with 13C-enriched carbonyls. e C-7–C-15 give these six signals. f C-8–C-15 give these seven signals. g C-9–C-15 give six signals. h C-10–C-15 give six signals. i C-4–C-6 give one signal. j C-12–C-15 give three signals. k C-4–C-7 give these four signals. l C-13–C-15 give these three signals. m C-4–C-9 give these four signals. n C-4–C-10 give these four signals. o C-4–C-11 give these five signals. p C-4–C-13 give these seven signals.

2.4.4.6. Monomethyl hexadecanedioate (5k). 85% yield. MS (CI, CH4) m/z 301.24 (MH+, 4.73%), 283.224 (MH+ H2O, 100%), 269.207 (MH+ MeOH, 99.31%); anal. HRMS (CI): calcd. (C17H33O4, MH+) 301.2379 obsd. 301.2395. 2.4.5. General procedure for preparation of acyl chloride monoester 6 Acyl chloride monoesters 6 were prepared in quantitative yields according to the procedure of Moreau et al. (2005) and minimally characterized by NMR. The acyl chlorides were stored under reduced pressure until used. The unlabeled analogs 6a, b and g are commercially available (Sigma–Aldrich). The doubly labeled analogs 6a, e and h were prepared but not characterized because of their sensitivity to moisture. 2.4.6. General procedure for preparation of keto esters 7 Keto esters 7 were prepared via the cadmium mediated coupling of acyl chlorides 6 with alkyl bromides 9, according to the procedure of Menger et al. (1989). The procedure was carried out in toluene (rather than benzene) and the compounds were purified by recrystallization from hexane. Partial spectral data for many of the analogs appear in the literature (Tulloch, 1977). The unlabeled analogs 7b, d, e–h and j are commercially available (Sigma–Aldrich). The major side product in this reaction is presumably the long chain alkane dimer 12, which was identified by its broad overlapping multiplets in 1H NMR absorbing at ca. 1–2 ppm. The long chain aliphatics are well known in the literature. 2.4.6.1. Methyl 4-oxooctadecanoate (7a). 44% yield, mp: 54–57
C. MS (DCICH4) m/z: 311.259 (MH+, 21.47%), 281.248 (MH+ CH3OH, 100%). HRMS (DCI CH4) m/z: calcd. (C19H35O3, M H+) 311.2586, obsd. 311.2592. FTIR: 2960 (s), 2880 (m), 1740 (m, CQO ester), 1710 (s, CQO ketone), 1470 (m), 1380 (m), 1230, 1172 (m, C O) cm1.

2.4.6.2. Methyl 4-oxooctadecanoate-1,4-13 C2 (13C-7a). mp: 54–57
C.

36% yield,

2.4.6.3. Methyl 6-oxooctadecanoate (7c). 49% yield, mp 43
C. MS (DCI, CH4) m/z 313.278 (MH+, 100%), 281.244 (MH+ MeOH, 100%), anal. HRMS (CI, CH4): calcd. (C19H37O3, MH+) 313.2770 obsd. 313.2783. FTIR 2960 (s), 2930 (s), 2880 and 2870 (m), 1740 (m, CQO ester), 1720 (m, CQOketone), 1650 (m), 1510 (w), 1460 (m), 1220 and 1168 (m, C O) cm1. 2.4.6.4. Methyl 8-oxooctadecanoate-1,8-13C2 (13C-7e). The product was obtained as an orange oil (0.598 g, 1.904 mmol, 100%) but resisted recrystallization from hexane. It was, therefore, used without further purification. 2.4.6.5. Methyl 1,12-13C-12-oxooctadecanoate (13C-7h). 89% yield, mp: 46–48
C. MS (DCICH4) m/z: 315.279 (MH+, 53.07%), 283.272 (MH+ CH3OH, 34.79%), 229.191 (MH+ C6H14, 100%). HRMS

Table 6 13 C NMR Shifts (ppm) for C-1 and C-7 in methyl 7-oxooctadecanoate (7d) in pure solvents and intercalated within DMPC liposomes.a Solvent

ET(30)b

Carbonyl C-7d

Carbonyl C-1

C6D6 Acetone-d6 CD3CN CD3OD DMPC liposomes

34.3 42.2 45.6 55.4

208.44 209.40 211.74 213.84 209.43 39.1

173.20 174.00 174.71 175.70 174.56 47.2

Calculated ETc :

a Experimental error is 0.05 ppm in chemical shift and c  1 kcal/mol in calculated ET (Cohen et al., 2008a). b kcal/mol at 25
C. c From Fig. 4. d See Supplementary material Fig. S2.

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M. Afri et al. / Chemistry and Physics of Lipids 184 (2014) 105–118

(DCI CH4) m/z: calcd. (12C1713C2H37O3, MH+) 315.2810, obsd. 315.2788.

299.2569. FTIR (KBr) 3061 (vbr, s, OH), 2926, 2845 (s,  CH2 ), 1703 (s, CQO) cm1.

2.4.6.6. Methyl 13-oxooctadecanoate (7i). 37% yield, mp: 43
C. MS (DCI, CH4) m/z 313.275 (MH+, 100%), 281.265 (MH+ MeOH, 6.35%), 197.071 ((CH2)11CO, 78.06%); anal. HRMS (CI): calcd. (C19H37O3, MH + ) 313.2743 obsd. 313.2754. FTIR (KBr) 2983 (w,  CH2 ), 2954 (m,  CH2 ), 2931, 2918 and 2849 (s,  CH2), 1738 and 1705 (s, CQO), 1470, 1438, 1415, 1380 and 1321 (m, CH), 1260, 1229, 1203 and 1176 (m, CO) cm1.

2.4.7.9. 12-Oxooctadecanoic acid (8h). 84% yield, mp: 110–112
C. MS (DCI CH4) m/z: 299.254 (MH+). HRMS (DCI CH4) m/z: calcd. (C18H35O3, MH+) 299.2586, obsd. 299.2542. FTIR (KBr): 3052 (vbr, s, OH), 2912, 2845 (s, CH2), 1696 (s, CQO) cm1.

2.4.6.7. Methyl 16-oxooctadecanoate (7k). 25% yield. MS (CI, CH4) m/z 313.272 (MH+, 34.73%), 283.217 ((MHEt), 100%), 281.249 (MH+MeOH, 61.89%); anal. HRMS (CI): calcd. (C19H37O3, MH+) 313.2743 obsd. 313.2724. 2.4.7. General procedure for preparation of keto acid 8 Keto acids 8 were prepared from the corresponding keto esters 7 by saponification according to the procedure of Menger et al. (1989). The products were crystalline. 2.4.7.1. 4-Oxooctadecanoic acid (8a). 90% yield, mp: 84
C. MS (DCI CH4) m/z: 299.258 (MH+, 23.36%), 281.239 (MH+ H2O, 100%). HRMS (CICH4) m/z: calcd. (C18H35O3, MH+) 299.2586, obsd. 299.2585. FTIR (KBr): 3405 (vbr, s, OH), 2919 (s), 2851 (m), 1703 (s, CQO) cm1. 2.4.7.2. 4-Oxooctadecanoic acid-1,4-13C2 (13C-8a). 90% yield, mp: 84
C. 5-Oxooctadecanoic acid (8b):89% yield, mp: 80
C; MS (EI+) m/z 298 (M+, 2.4%), 211 (M+ (CH2)3COOH, 15.6%), 84 (C6H12, 100%). 2.4.7.3. 6-Oxooctadecanoic acid (8c). 60% yield, mp 84
C. MS (DCI, CH4) m/z 299.259 (MH+, 9.8%), 281.243 (MH+ H2O, 100%). FTIR 3599 (vbr, s, OH), 2976 (m), 2917 (s), 2846 (m), 1715 and 1688 (s, CQO), 1473 (m), 1481 (w), 1260 and 1105 (m, C O) cm1. 2.4.7.4. 7-Oxooctadecanoic acid (8d). 13% yield, mp: 78
C. MS (DCI, CH4) m/z 299.259 (MH+, 48.19%), 281.249 (MH+ H2O, 100%), 158.095 ((CH2)10CH3, 18.84%); anal. HRMS (CI): calcd. (C18H34O3, MH+) 299.2586 obsd. 299.2587. FTIR (KBr) 3105 (vbr, s, OH), 2954, 2928, 2917, 2848 (s,  CH2 ), 1715, 1699 (s, CQO), 1470, 1419 and 1382 (m, CH), 1250 and 1106 (m, C O) cm1. 2.4.7.5. 8-Oxooctadecanoic acid (8e). 82% yield, mp: 79
C. MS (DCI CH4) m/z: 299.258 (MH+, 13.22%), 281.261 (MH+ H2O, 100%), 157.084 (MH+ C10H22, 88.61%). HRMS (DCI CH4) m/z: calcd. (C18H35O3, MH+) 299.2586, obsd. 299.2576. FTIR: 3737 (vbr, m, OH), 2969, 2939 (s,  CH2), 2882 (m), 1749, 1737 (s, CQO), 1541, 1457, 1376 (w), 1217 (w, C O) cm1. 2.4.7.6. 8-Oxooctadecanoic acid-1,8-13C2 (13C-8e). 62% yield, mp: 79
C. MS (DCI CH4) m/z: 283.257 (MH+ OH, 0.79%). HRMS (DCI CH4) m/z: calcd. (12C1613C2H33O2, MH+ OH) 283.2548, obsd. 283.2573. 2.4.7.7. 9-Oxooctadecanoic acid (8f). 95% yield, mp: 74
C. MS (DCI, CH4) m/z 298.2 (MH+, 7.22%), 281.247 (MH+ H2O, 100%), 155.108 (CO(CH2)7CO, 21.79%), 125.063 ((CH2)7CO, 12.8%), 98.047 ((CH2)7, 21.74%). anal. HRMS (CI): calcd. (C18H34O3, M+) 298.2508 obsd. 298.2437. FTIR (KBr) 3200–3014 (vbr, s, OH), 2955, 2929, 2919 and 2848 (s, CH2 ), 1717, 1699 (s, CQO), 1470, 1437, 1380, 1306 and 1332 (m, C H), 1106 (w, C O) cm1. 2.4.7.8. 10-Oxooctadecanoic acid (8g). 26% yield, mp: 80
C. MS (DCI CH4) m/z: 299.257 (MH+, 18.1%), 281.251 (MH+ H2O, 100%). HRMS (DCI CH4) m/z: calcd. (C18H35O3, MH+) 299.2586, obsd.

2.4.7.10. 12-Oxooctadecanoic acid-1,12-13C2 (13C-8h). 87% yield, mp: 110–112
C. MS (DCI CH4) m/z: 301.262 (MH+, 8.62%), 284.252 (MH+OH, 18.44%), 283.251 (MH+ H2O, 100%), 215.153 (MH+ C6H14, 40.82%). HRMS (DCI CH4) m/z: calcd. (12C1613C2H35O3, MH+) 301.2653, obsd. 301.2617. 2.4.7.11. 13-Oxooctadecanoic acid (8i). 60% yield, mp: 82
C. MS (DCI, CH4) m/z 299.25 (MH+, 15.38%), 281.24 (MH+ H2O, 100%), 99.052 (CH3(CH2)4CO, 12.13%), 71.07 (CH3(CH2)4, 6.5%); anal. HRMS (CI): calcd. (C18H35O3, MH+) 299.2586 obsd. 299.2539. FTIR (KBr) 3115 (vbr, s, OH), 2954, 2930, 2916 and 2849 (s,  CH2 ), 1699 (s, CQO), 1469, 1440, 1418 and 1317 (m, CH), 1266, 1236, 1214 (w, C O) cm1. 2.4.7.12. 14-Oxooctadecanoic acid (8j). 35% yield, mp: 82
C. MS (CI, CH4) m/z 299.260 (MH+, 7.71%), 281.248 (MH+ H2O, 100%); anal. HRMS (CI): calcd. (C18H35O3, MH+) 299.2586 obsd 299.2596. FTIR (KBr) 3061 (vbr, s, OH), 2912, 2851(s, CH2 ), 1703 (s, CQO) cm1. 2.4.7.13. 16-Oxooctadecanoic acid (8k). 40% yield, mp: 84
C. MS (CI, CH4) m/z 299.258 (MH+, 5.45%), 281.252 (MH+ H2O, 100%); anal. HRMS (CI): calcd. (C18H35O3, MH+) 299.2586 obsd. 299.2583. FTIR (KBr) 3074 (vbr, s, OH), 2919, 2851 (s, CH2 ), 1703 (s, CQ) cm1. 2.5. NMR conditions for preparing correlation graphs and liposomal samples For the correlation graph, samples (ca. 10 mg) of ketoesters 7 or ketoacid 8 were dissolved in 0.6 mL of pure solvent to give a final concentration of 0.05 M. This is approximately the same concentration as that of the intercalants within the liposomal solution. The samples in pure solvents were scanned for 20 min. or 1000 scans. Liposomal samples were prepared as described above in Section 2.3 and were scanned overnight (40,000 scans). D2O served as the internal lock. 3. Results and discussion 3.1. General synthesis of keto esters 7 and keto acids 8 As pointed out in the introduction, the first goal of this research was to synthesize two related families of compounds containing two carbonyl groups, long chain fatty keto esters 7 and keto acids 8. Each of these families would be comprised of the same number of carbons and functional groups, with the ester or acid carbonyls presumably anchored in the polar region of the membrane. The only difference between the various members in each set would be the location of the second ketone carbonyl along the fatty acid chain. As outlined in Fig. 3 above, the first step in the synthesis of these families was the quantitative conversion of dibromoalkanes 1 to the corresponding dicyanoalkane derivatives 2 using KCN, followed by basic hydrolysis of 2 to diacids 3. Monoesterification of 3 to give monoacid monoester 5 proved to be a low-yield approach. More effective was the initial diesterification of diacid 3 to diesters 4 followed by monohydrolysis to the desired acid esters

M. Afri et al. / Chemistry and Physics of Lipids 184 (2014) 105–118

3.2. Correlation graphs of keto esters 7 and keto acids 8 With keto esters 7 and keto acids 8 now in hand, we proceeded to measure the chemical shifts of the various carbons in each different derivatives in solvents of various ET(30) polarities. As expected, based on extensive previous experience, only the polarizable carbonyls were substantially and conasistently responsive to polarity changes. A 13C NMR chemical shift–solvent polarity correlation graph was created for each homolog in each family, and the correlation coefficient R2 was determined to confirm reliability and consistency of our results. As noted in our introductory comments, these correlation graphs make it possible to determine the polarity of the microenvironment experienced by a specific carbonyl based on the observed chemical shift. From this, the carbonyl’s qualitative depth within the liposome can be estimated using the correlation graph. This process will be exemplified by describing the formation and use of a correlation graph of methyl 7-oxooctadecanoate (7d; Fig. 3) and the corresponding keto acid 7-oxooctadecanoic acid (8d). Ketoeser 7d was dissolved in four solvents (ET(30) values in kcal/mol from Reichardt, 1994): benzene (34.3), acetone (42.2), acetonitrile (45.6) and methanol (55.5). The 13C NMR chemical shifts for each carbonyl carbon were plotted as a function of solvent

C-7 (keto) 2

R = 0.94

Liposome benzene

acetone acetonitrile

methanol

C NMR Chemical shift (ppm)

EtPenOH dMePenOH

MeBuOH

190

C-1 (acid)

2

2

R = 0.94

180

2

180

R = 0.98

R = 0.98

30

35

40

45

50

55

60

E T(30) (kcal/mol) Fig. 4. The change in 13C NMR chemical shift of carbonyls C-1 and C-7 in methyl 7oxooctadecanoate (7d) as a function of the ET(30) polarity of pure solvents. The hollow symbols represent the corresponding liposome value. (Errors: ca. 1.0 kcal/ mol in the ET(30) value and 0.05 ppm in chemical shift. Errors are smaller than the data points.)

2

R = 0.04

170 160 35

30

40

45 50 E T(30) (kcal/mol)

55

60

Fig. 5. The change in 13C NMR chemical shift of carbonyls C-1 and C-7 in 7oxooctadecanoic acid (8d) as a function of the ET(30) polarity of pure solvents. The hollow symbols represent the corresponding liposome value. (Errors are as detailed in Fig. 4.)

polarity. As expected, the correlation graph of keto ester 7d (Table 6 and Fig. 4) revealed a very good direct correlation between the polarity of the solvent and the chemical shifts of the ketone (R2 = 0.94) and the ester carbonyls (R2 = 0.98). These results repeat themselves closely for all eleven members of keto ester family 7 synthesized (7a–k). Turning now to the keto acids 8, and 7-oxooctadecanoic acid (8d) is emblematic of the whole family. When drawing up our initial correlation graph (not shown) in the abovementioned four solvents (benzene, acetone, acetonitrile and methanol), we observed an excellent correlation (R2 = 0.98) between the solvent polarity and the chemical shifts of the ketone carbonyl. However, the corresponding line for the acid carbonyls gave a very poor correlation coefficient value of R2 = 0.11. For this reason, a new correlation graph was prepared based on a much wider range of eight solvents (abbreviation in Fig. 5; ET(30) values in kcal/mol from Reichardt, 1994): benzene (34.3), 3-ethylpentanol (EtPenOH; 38.5), 2,4-dimethyl-3-pentanol (dMePenOH; 40.1), 2methyl-2-butanol (MeBuOH; 41.0), acetone (42.2), acetonitrile (MeCN; 45.6), ethanol (EtOH; 51.9), and methanol (MeOH; 55.5). The new correlation graph obtained (Fig. 5) again showed an excellent correlation for the keto group (R2 = 0.96) but a poor correlation coefficient of R2 = 0.04 for the acid carbonyl (see the dashed line in Fig. 5). Further examination of this latter curve reveals that there is a break in this line at ca. 43 kcal/mol (just above acetone). As one proceeds to more polar solvents from this point (increasing ET(30) values), one observes the expected direct relationship between

R C OH

O R C OH

Increasing solvent polarity Increases acyl carbon chemical shift O

160

MeOH

acetone

O C-1 (ester)

2

R =0.96

EtOH

Liposome

200

C-7 (keto)

MeCN

benzene

210

200

13

C NMR Chemical Shift (ppm)

220

220

13

5. Esterification was accomplished by ultra-sonication in acidified methanol, which has various advantages: (a) simplicity; (b) the absence of hazardous reagents; (c) high yields; (d) the absence of side products; and (e) relatively short reaction times (generally, a few hours). The monohydrolysis of diesters 4 was mediated by Ba (OH)2, yielding the desired monoester monoacids 5. The shorter analogs, monomethyl succinate (5a) and monomethyl glutarate (5b), were conveniently prepared via the methanolysis of succinic (13a) and glutaric (13b) anhydrides, respectively. Monoester monoacids 5 were quantitatively converted with thionyl chloride to the acyl chlorides 6, which were alkylated in turn with the appropriate dialkylcadmium reagent 11 to give the 18-carbon methyl keto esters 7 accompanied by variable amounts of the Wurtz coupling product 12. The keto ester underwent base catalyzed hydrolysis to the desired keto acid 8. By beginning with 13 C enriched K13CN (Fig. 3, step A) or doubly labeled succinic anhydride (13a, step J), one could prepare the corresponding labeled dicarbonyls compounds 7 and 8, homologs a, e, and h. The 1H and 13C NMR spectral data of compounds 2–8 appear in Tables 1–5.

113

R C OH

O R C O

+H

Increasing solvent polarity Decreases acyl carbon chemical shift Fig. 6. Opposing effects of solvent polarity on the acid carbonyl chemical shift.

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M. Afri et al. / Chemistry and Physics of Lipids 184 (2014) 105–118

Table 7 Chemical shifts (ppm) and corresponding ET(30) values (kcal/mol)a of carbons C1 and Cn of keto esters 7a–k intercalated within the DMPC liposomes. Cmpd. (n)

7a(n = 4) 7b(n = 5) 7c(n = 6) 7d (n = 7) 7e(n = 8) 7f (n = 9) 7g(n = 10) 7h(n = 12) 7i(n = 13) 7j(n = 14) 7k(n = 16) a b c

d C1 (ppm) c

n.d. 174.38 174.6 174.57 174.4 174.31 173.8 173.8 173.83 n.d.c n.d.c

ET(30) C1 – 45.7 45.2 45.8 43.5 41 40.3 37.5 37.2 – –

d Cn (ppm)

ET(30)a Cn

Dd(C1  Cn)Calc.

208.92 209.83 209.85 209.44 209.1 208.73 208.4 208.1 208.15 208.7 206.8

40.5 41 40.9 38.1 36.1 33.2 32.2 31.8 31.6 30.8 31.3

3.75 5 6.25 7.5 8.75 10 11.25 13.75 15 16.25 18.75

(Å)b

These ET(30) values are calculated from the correlation curve and have an error of ca. 1.0 kcal/mol (Cohen et al., 2008a). Calculated with PCMODEL 7.50.00 Software (Bloomington, Indiana) using an MMX force field. n.d = not determined (Section 3.3.1).

chemical shift and solvent polarity. Thus, as solvent polarity rises so does the chemical shift of the carboxy carbonyl, with an excellent correlation coefficient of R2 = 0.98. However, below the break, the trend is inverted: as solvent polarity drops, the chemical shift of the carboxy carbonyl also rises – again with an excellent correlation coefficient of R2 = 0.94. A similar correlation graph line-break has been previously observed in other carboxylic acid systems (Weitman et al., 2001; Afri et al., 2004). In the latter work it was thought that the break had to do with whether or not the solvent was protic. However, from the present study, it would seem clear that this is not the case, since there are protic and non-protic solvents on both sides of the break. Rather, it is now believed that the problematics of the carboxylic acid system can be attributed to the presence of two competing phenomena (Fig. 6). As already noted in the introduction, as the solvent polarity increases, the charge-separated zwitterionic resonance form of the carbonyl makes a concomitantly greater contribution. As a result, the NMR chemical shift of the carbonyl carbon rises. However, in the case of carboxylic acids, there is an additional contravening factor at play: the equilibrium between the associated (COOH) and dissociated forms (COO) of the acid moiety. As the solvent polarity increases, there is concomitantly greater dissociation of the carboxylic acids with a build up of negative charge on the carboxylate oxygen (Reichardt and Welton, 2011 p. 109). This build-up spills over to the acid carbonyl carbon, whose positive charge is increasingly neutralized by the electron rich oxyanion. This in turn results in a lowering of the chemical shift of the acid carbonyl. Clearly, the amount of positive charge on the acid carbonyl carbon at each polarity value will depend on the relative contributions of each of these factors. At higher ET(30) values, the polarization of the carbonyl is the predominant factor and, hence,

the chemical shift-polarity correlation is a positive one. However, at low polarity, polarization of the carbonyl becomes minimal, and the determinant factor becomes the amount of acid dissociation. It is clear, then, that no single line equation is appropriate for the acid carbonyl correlation curve over the whole range of polarity values. However, truth be told, a complete correlation graph is not required. This is because, the acid groups in all the derivatives of 8 lie in the upper polar region of the liposomal membrane (vide infra), not far from the interface. Hence, the correlation curve obtained using just the polar solvents (acetone, acetonitrile, ethanol and methanol) can be used – for which region, as noted above, the correlation coefficient is excellent (R2 = 0.98). 3.3. Intercalation of keto esters 7 and keto acids 8 into liposomes With suitable correlation graphs in hand, the next step was to intercalate the substrates within DMPC liposomes, as previously described (Frimer et al., 1996; Afri et al., 2002, 2004a,b; Bronshtein et al., 2004c; Cohen et al., 2008a,b,c; Bodner et al., 2010), and measure the 13C NMR chemical shifts for the acyloxy carbonyl carbons. The data from the respective correlation graphs allowed us to calculate the ET(30) polarity value corresponding to each chemical shift observed for the carbonyl carbons of keto esters 7 and keto acids 8 intercalated in the liposome (see Tables 7 and 8). In addition these tables contain Dd(C1  Cn)Calc, which is the aerial gap difference (in Å) between the upper acid or ester carbonyl (C1) and the deeper lying ketone carbonyl (Cn) – calculated using PCMODEL software. At this juncture, it would be appropriate to discuss the use of PCMODEL MMX calculations for measuring distances in these long-chain systems. One could well argue that static molecular models of these long dicarbonyl fatty acid chains and

Table 8 Chemical shifts (ppm) and corresponding ET(30) values (kcal/mol) of carbons C1 and Cn of keto acids 8a–k intercalated within the DMPC liposomes.a Compd. (n)

8a (n = 4) 8b(n = 5) 8c(n = 6) 8d(n = 7) 8e(n = 8) 8f(n = 9) 8g(n = 10) 8h(n = 12) 8i(n = 13) 8j(n = 14) 8k(n = 16) a

See notes to Table 7.

d C1 (ppm) 176.76 177.17 177.56 177.21 177 176.95 177 176.71 176.6 176.69 176.92

ET(30) C1

d Cn (ppm)

56.8 55.5 56.7 55.6 51.9 53.5 51.3 51.1 50.7 51.2 52.3

210.35 210.65 210.53 209.51 209.2 208.41 209.2 207.51 207.6 207.38 208.42

ET(30)a Cn

Dd(C1  Cn)Calc.

42.7 43.6 42.6 37.5 38.7 34.2 33.7 30 33 29.3 33.3

3.75 5 6.25 7.5 8.75 10 11.25 13.75 15 16.25 18.75

(Å)

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phospholipids generated by PCMODEL MMX calculations are an oversimplification of reality and unlikely to yield a proper model of a fluid bilayer. This is because lipid hydrocarbon chains in the fluid bilayer phase have a certain probability of trans/gauche isomerization. Furthermore, lipid molecules undergo rapid wobble. Both effects are temperature dependent and they reduce bilayer thickness and increase area per molecule. Noteworthy, however, are recent diffraction studies by Ku9 cerka et al. (Ku9 cerka et al., 2011) which have shown that the aforementioned effects are relatively small, 4% over a 20
range, and hence should certainly not have any dramatic affect on the preliminary results reported herein. Not surprisingly, these studies also show that in saturated fatty acid chains trans/gauche isomerization is substantially tempered by attractive chain–chain van der Waals interactions which increase with increasing chain length. Most importantly, PCMODEL MMX calculations estimated the length of one-half of a bilayer – from the choline nitrogen down to the midplane – as ca. 22 Å. This corresponds nicely to bilayer thickness values of ca. 44 Å reported for DMPC liposomes (Ku9 cerka et al., 2004, 2005), and other experimental data reviewed by Cohen et al. (2008a). All this strongly confirms the reliability of the MMX calculations. There is, however, one assumption which may well deserve further investigation, and that is, that the thickness of the DMPC lipid bilayer does not change dramatically upon the intercalation of intercalants 7 and 8.

value for the ester carbonyl head group holds steady for keto esters 7b–d (n = 5–7) at ca. 45.5 kcal/mol. But then it drops some 8.5 units down to 37 kcal/mol for keto ester 7i (n = 13), spanning much of the non-polar region of the liposome. From previous studies (Cohen et al., 2008a), it is known that the two glycerol carbonyls in the DMPC liposome are situated at a polarity of ca. 50–52 kcal/mol. Thus, an ET(30) value of 45.7 kcal/mol places keto ester 7b somewhat below the glycerol ester carbonyls of the DMPC liposomes toward the top of the non-polar region. By contrast, 37 kcal/mol places keto ester 7i substantially down along the 14-carbon DMPC lipid chain. Perforce, one must conclude that the C1  Cn proximity has a dramatic affect on the “buoyancy” of the upper part of the keto ester chain. Turning now to the ketone carbonyl carbons, as expected their chemical shifts are substantially lower, spanning some 9 ET units from 40.5 kcal/mole in compound 7a down to 31.3 kcal/mol in keto ester 7k. The former value indicates that the ketone of 4-keto ester 7a lies toward the upper portion of the lipid fatty acid chain. Regarding the lower value of 31.3 kcal/mol in keto ester 7k, this is well along the fatty acid chain considering that 31 kcal/mol is the lowest possible value on the ET(30) scale. Indeed, the ketone carbonyl ET(30) values of Table 7 suggest that, as this moiety penetrates deeper into the bilayer, the polarity it senses approaches the minimum value 31 kcal/mol asymptotically – as would be expected for an exponential function. We will return to this point momentarily.

3.3.1. The ET(30) polarity values of keto esters 7 We begin with several preliminary comments. Firstly, a quick initial survey of the data in Table 7 for keto esters 7 reveals that no chemical shifts were determined for the ester carbonyl of keto esters 7a, j and k. This is because the DMPC glycerol esters of the liposome have very large and broad absorptions at ca. 173–174 ppm, which sometimes obscure other ester peaks falling in the same range. Secondly, in interpreting the results of Table 7, it seems appropriate to assume that keto esters 7a–k lie essentially vertically in the liposomes, parallel to the liposomal lipids, based on the following reasoning. As mentioned above, there exists a polarity gradient as one moves deeper into the lipid bilayer – ranging from that of water (ET(30) = 63 kcal/mol) at the water–lipid interface down to hexane (ET(30) = 31 kcal/mol) deep in the non-polar lipid slab. Different molecules in the lipid bilayer feel different polarities in accordance to their location and orientation. The keto esters 7a–k are amphiphilic, having dissimilar hydrophobic and hydrophilic regions. When intercalated into the liposome, the acyloxy moiety of the ester head group is expected to rise toward a rather polar region within the bilayer. On the other hand, the long fatty acid tail is expected to pull the intercalant toward the more hydrophobic lipid slab, allowing for greater van der Waals interactions with the long-chain lipid tails of the liposome. As a result, keto esters 7a–k are expected to lie vertically in the liposomes, parallel to the lipid chains and perpendicular to the water/lipid interface. Considering that all the keto ester derivatives 7a–k have the same number of carbons, and differ only in the location of the ketone group, it would have been expected that all the ester carbonyls would be anchored at approximately the same value. The ET(30) value for the ketone carbonyls would be expected to decrease as the “n” value increases and the ketone moves down the chain approaching the lipid slab. This, in turn, should render the construction of a chemical ruler quite trivial. Much to our frustration, the behavior and observed values for keto esters 7 given in Table 7 are not as predicted. In particular, the ester carbonyls are not anchored near the lipid–water interface, nor are they even all located at the same depth. Indeed, the ET(30)

3.3.2. ET(30) polarity values of keto acids 8 The results for keto acids 8 appear in Table 8. Unlike the ester derivatives, the 13C chemical shifts of all the acyloxy carbons were detected, because the carbonyl peaks fall downfield (>176.6 ppm) from the liposomal DMPC ester carbonyls (173–174 ppm). The results indicate that as expected the carboxylate carbonyls are located in the polar region of the liposome; however, they are not anchored at the water–lipid interface (at ca. 63 kcal/mol) or even at some other very polar value, as generally assumed in the literature (Bronshtein et al., 2004). Rather, they range over ET(30) polarity values of 51–57 kcal/mol. These results are not inconsistent with previous work from our lab (Cohen et al., 2008a) which has shown that the interface region between the lipid chains and the aqueous media is actually a broad area, which spans about 12 ET(30) units (63–51 kcal/mol). This area extends from the charged choline group, past the zwitterionic phosphate moiety, down to the glycerol ester carbonyls. Some ions and small polar molecules such as water have been shown to readily penetrate into this area. (Gamliel et al., 2008). Interestingly, the ET values for the acid carbonyls remains rather constant at ca. 56 kcal/mol for the shorter derivatives 8a–d (n = 4–7). However, this is followed by a sudden drop of 4 ET units to 52 kcal/mol for 8e (n = 8), and the polarity value again remains essentially constant at 51–52 kcal/mol for the remaining derivatives 8e–k (n = 8–16). Interestingly, as in keto esters 7, the breaking point for the acid carbonyl is in-between n = 7 (8d) and n = 8 (8e). As before, the drop is presumably related to the relative hydrophilicity of the upper portion of the keto acid. The results for the ketone carbonyls of keto acids 8 are similar mutatis mutandis to those observed in the keto ester 7 system. The polarity of the keto-carbonyl carbon decreases as the distance from the polar head group increase. Here too, there is a sudden sharp drop in ET(30) value in-between n = 7 (8d) and n = 8 (8e). But again this is not surprising assuming that the chain is lying perpendicular to the interface and the head and tail portions of the chain move together. Interestingly, in the shallow region (4  n  6) the average polarity of the ketone carbonyl is 43.1 kcal/mol, while in the deeper area (12  n  16) the average polarity is ca. 33 kcal/mol, placing it deep within the DMPC lipid slab.

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This process was repeated iteratively several times, until obtaining the best fit required no further corrections. Fig. 7 shows the “best fit” curve obtained for the data points of keto esters 7 and keto acids 8. The exponential curve gives us two mathematical relationships between PD and the corresponding ET(30) value, as shown in Eqs. (2) and (3).

Fig. 7. ET(30) values sensed by the keto ester 7 and keto acid 8 carbonyls as a function of penetration depth (PD).

3.4. The change in ET(30) as a function of penetration depth (PD) For the purpose of constructing a molecular ruler, it is important to determine how the ET value changes as a function of the penetration depth (PD) within the liposome. As previously explained, the original plan was to develop a molecular ruler using families of isomeric compounds containing two polarizable functional groups; the upper moiety would be anchored in or at the interphase, while the lower one would progress gradually down the chain. Since the ester and acid functionalities of derivatives 7 and 8 were not anchored as hoped for, a less straight-forward more sophisticated approach for gleaning the desired information was necessary. Because the lipids comprising the liposomal bilayer are amphiphilic, two rather different areas of polarity, one polar and the other apolar, would be a priori expected. As noted in our introductory comments, a gradient of polarities was indeed observed – ranging from that of water (ET(30) = 63 kcal/mol) at the water–lipid interface, down to hexane (ET(30) = 31 kcal/mol) deep in the non-polar lipid slab. Sharp transitions in polarity do not normally exist in nature on the bulk scale, unless one is dealing with different sides of a membrane or other partition. This corresponds best to an exponential function, which has a penetration depth (PD) (in Å) zero at ca. ET(30) = 63 kcal/mol at one end, and approaches the minimum ET(30) value at ca. 31 kcal/ mol asymptotically at the other. To calculate this exponential function, as a first approximation, all the head group carbonyls (C1) of the keto esters 7 and keto acids 8 were lined up on the Y-axis according to their determined ET(30) values (see Tables 7 and 8) at zero penetration (X-axis). The tail group carbonyls were then placed on the graph based on their determined ET(30) values and an approximate PD, assuming that the latter value is governed by Eq. (1): PDCn ¼ PDC1 þ DdðC1  CnÞ

(1)

where Dd(C1  Cn)Calc is the PCMODEL calculated distance from C1 to Cn in Å, listed in Tables 7 and 8. The curve drawing program was then asked to find the exponential curve that would give the “best fit.” Once this was done, the head groups were then moved to the curve and the “best fit” process repeated. It should noted that in the case of the keto acids, since the headgroup carbonyls lie at the beginning of the curve where the slope is rather steep, only small corrections were required. For the keto esters, however, the initial correction was a bit more substantial. Clearly, a revision of the initial PD value of the C1 headgroup carbonyls had a concomitant effect on the location of the tailgroup carbonyls, based on Eq. (1).

ET ð30Þ ¼ 31:5 þ 31:5  expð0:137PDÞ

(2)

  31:5 PD ¼ 7:30  ln ET ð30Þ  31:5

(3)

From a practical perspective, using Eq. (3) to calculate PD values from the corresponding ET(30) numbers works well (i.e., gives good results with only small errors) only in the more polar area of the bilayer (ET(30) 0 37). However, as one moves to the very lipophilic areas of the liposomal bilayer – as is the case for the PD of the Cn very long chain keto esters and keto acid and, a slight error in ET leads to a very large error in the calculated PD. As a result, a more reliable means for determining the PD of Cn, is to calculate the PD of the headgroup C1 using Eq. (3), and then adding to it the PCMODEL calculated distance from C1 to Cn, Dd(C1  Cn)Calc. This in effect is Eq. (1). Table 9 lists the PDCalc of carbons C1 keto esters 7 and 8, calculated from the corresponding ET(30)exp values of Tables 7 and 8 using Eq. (3). The PDCalc of carbons Cn was calculated using Eq. (1), as just described. It should be noted that since keto esters 7a, j and k lack points for C1, they were not included in Fig. 7 and Table 9. Also not included are keto acids 8h and j which yield ET (30) ET values for the ketone carbonyl which are below 31 kcal/mol, the minimal value possible. The curve in Fig. 7 raises several issues which require clarification. Firstly, while the data points for keto esters 7 and keto acids 8 all lie in close proximity to the “best fit” curve, the keto ester points as a rule tend to lie on or above the curve, while those of the keto acid seem to lie on or somewhat below. This is presumably related to the fact that the substrate (intercalant) is present in a substrate:lipid molar ratio of 1:5. As a result, the polarity sensed by the carbonyls are to some small extent a result of the substrate itself, and not merely the solvent. Having a large number of points from different substrates should allow us to cancel this effect out to some extent. Table 9 ET(30) value (kcal/mol) and calculated penetration depth (PD in Å)of carbons C1 and Cn of keto esters 7 and 8 intercalated within the DMPC liposomes. Cmpd. (n)

8a (n = 4) 8b(n = 5) 8c(n = 6) 8d(n = 7) 8e(n = 8) 8f(n = 9) 8g(n = 10) 8i(n = 13) 8k(n = 16) 7b(n = 5) 7c(n = 6) 7d (n = 7) 7e(n = 8) 7f (n = 9) 7g(n = 10) 7h(n = 12) 7i(n = 13) a b c d

ET(30) C1

PD C1

PD Cn

ET(30) Cn

Exp.a

Calc.b

Calc.c

Exp.a

Calc.d

56.8 55.5 56.7 55.6 51.9 53.5 51.3 50.7 52.3 45.7 45.2 45.8 43.5 41 40.3 37.5 37.2

1.64 2.03 1.67 2 3.25 2.68 3.47 3.7 3.1 5.95 6.22 5.9 7.21 8.95 9.52 12.38 12.76

5.39 7.03 7.92 9.5 12 12.68 14.72 18.7 21.85 10.95 12.47 13.4 15.96 18.95 20.77 26.13 27.76

42.7 43.6 42.6 37.5 38.7 34.2 33.7 33 33.3 41 40.9 38.1 36.1 33.2 32.2 31.8 31.6

46.6 43.5 42.1 40.1 37.6 37 35.7 33.9 33.1 38.5 37.2 36.5 35 33.8 33.3 32.4 32.2

From Tables 7 and 8. From Eq. (3). From Eq. (1), where PD C1 was calculated from Eq. (3). From Eq. (2). The standard deviation for 17 points is 1.90.

M. Afri et al. / Chemistry and Physics of Lipids 184 (2014) 105–118

Secondly, the thickness of one-half of the DMPC bilayer is estimated to be ca. 22–23 Å (Cohen et al., 2008a). According to the curve in Fig. 6 this occurs at an ET(30) value of about 32–33 kcal/mol Yet the curve places the tail carbonyl of the longest keto esters 7h (n = 12) and 7i (n = 13) at ca. 27 Å. This data can be readily resolved in one of two ways. The first is that the tail carbons cross the midplane and begin to penetrate the complementary half of the bilayer. In this lower region of the second half of the lipid layer, the ET(30) polarity is also expected to be ca. 32–33 kcal/mol. An alternative suggestion is that the long tails situate themselves in the midplane. This latter approach has been widely used to describe the location of the ubiquinone tails, or even rings, within the bilayer (see: Afri et al., 2004a for a review). The final issue relates to the completeness of our results. Combining keto ester 7 and keto acids 8 together to construct the molecular ruler curve in Fig. 7 was a serendipitous choice. The former family supplies a good number of points in the apolar section of the curve, while the latter affords many points for the more polar region. Although the points nicely fit the curve with a standard deviation of 1.9 kcal/mol, experimental points are lacking for the region between ET(30) values of 51 and 45.5 kcal/mol. It would, therefore, seem appropriate to add more points in this region of the curve. To this end, the companion paper describes the preparation of a series of ketophosphatidylcholines, whose ester carbonyls fall exactly in this region (Afri et al., 2014). 4. Conclusions We had originally hoped to develop a molecular ruler using families of isomeric compounds containing two polarizable functional groups, where the upper moiety was anchored high in the bilayer, and the lower group would progress gradually down then chain. Although keto ester and keto acid derivatives 7 and 8 were not anchored as originally thought, use of an iterative best fit analysis of the data points obtained allows one to prepare an exponential curve. This curve provides substantial insight into the correlation between ET(30) micropolarity and penetration depth into the liposomal bilayer. Still lacking for the curve are data points in the ET(30) range of 51–45.5 kcal/mol. Once suitable substrates are prepared, we should be closer to a molecular ruler for testing and application. Transparency document The Transparency document associated with this article can be found in the online version. Acknowledgements The kind and generous support of the Israel Science Foundation (Grants Number 327/02 and 437/06) – founded by The Israel Academy of Sciences and Humanities, and The Ethel and David Resnick Chair in Active Oxygen Chemistry is gratefully acknowleged. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.chemphyslip.2014.07.007.

References Afri, M., Alexenberg, C., Aped, P., Bodner, E., Cohen, S., Eigenberg, M., Eliyahu, S., Harel, Y., Naqqash, M.E., Porat, H., Frimer, A.A., 2014. NMR-based molecular ruler for determining the depth of intercalants within the lipid bilayer. Part IV: Studies on KetoPhospholipids (Companion paper).

117

Afri, M., Ehrenberg, B., Talmon, Y., Schmidt, J., Cohen, Y., Frimer, A.A., 2004a. Active oxygen chemistry within the liposomal bilayer. Part III: locating vitamin E, ubiquinol and ubiquinone and their derivatives in the lipid bilayer. Chem. Phys. Lipids 131, 107–121. Afri, M., Frimer, A.A., Cohen, Y., 2004b. Active oxygen chemistry within the liposomal bilayer Part IV: locating 20 ,70 -dichlorofluorescein (DCF), 20 ,70 -dichlorodihydrofluorescein (DCFH) and 20 ,70 -dichlorodihydrofluorescein diacetate (DCFH-DA) in the lipid bilayer. Chem. Phys. Lipids 131, 123–133. Afri, M., Gottlieb, H.E., Frimer, A.A., 2002. Superoxide organic chemistry within the liposomal bilayer. Part II: a correlation between location and chemistry. Free Radic. Biol. Med. 32, 605–618. Afri, M., Naqqash, M.E., Frimer, A.A., 2011. Using fluorescence to locate intercalants within the lipid bilayer of liposomes, bioliposomes and erythrocyte ghosts. Chem. Phys. Lipids 164, 759–765. Altenbach, C., Greenhalgh, D.A., Khorana, H.G., Hubbell, W.L., 1994. A collision gradient method to determine the immersion depth of nitroxides in lipid bilayers: application to spin-labeled mutants of bacteriorhodopsin. Proc. Natl. Acad. Sci. U. S. A. 91, 1667–1671. Aussenac, F., Laguerre, M., Schmitter, J.-M., Dufourc, E.J., 2003. Detailed structure and dynamics of bicelle phospholipids using selectively and per-deuterated labels: a 2H NMR and molecular mechanics study. Langmuir 19, 10468–10479. Bodner, E., Afri, M., Frimer, A.A., 2010. Determining radical penetration into membranes using ESR splitting constants. Free Radic. Biol. Med. 49, 427–436. Bronshtein, I., Afri, M., Weitman, H., Frimer, A.A., Smith, K.M., Ehrenberg, B., 2004. Porphyrin depth in lipid bilayers as determined by iodide and parallax fluorescence quenching methods and its effect on photosensitizing efficiency. Biophys. J. 87, 1155–1164. Caputo, G.A., London, E., 2003. Using a novel dual fluorescence quenching assay for measurement of tryptophan depth within lipid bilayers to determine hydrophobic a-helix locations within membranes. Biochemistry 42, 3265–3274. Cohen, Y., Afri, M., Frimer, A.A., 2008a. NMR-based molecular ruler for determining the depth of intercalants within the lipid bilayer. Part II: quantitative values. Chem. Phys. Lipids 155, 114–119. Cohen, Y., Afri, M., Frimer, A.A., 2008b. Aggregate formation in the intercalation of long chain fatty acid esters into liposomes. Chem. Phys. Lipids 155, 120–125. Cohen, Y., Bodner, E., Richman, M., Frimer, A.A., 2008c. NMR-based molecular ruler for determining the depth of intercalants within the lipid bilayer. Part I: discovering the guidelines. Chem. Phys. Lipids 155, 98–113. Cohen, Y., Weitman, H., Afri, M., Yanus, R., Rudnick, S., Talmon, Y., Schmidt, J., Aped, P., Shatz, S., Ehrenberg, B., Frimer, A.A., 2012. The effect of intercalants on the host liposome. J. Liposome Res. 22, 306–318. Evanics, F., Hwang, P.M., Cheng, Y., Kay, L.E., Prosser, R.S., 2006. Topology of an outermembrane enzyme: measuring oxygen and water contacts in solution NMR studies of PagP. J. Am. Chem. Soc. 128, 8256–8264. Fanucci, G.E., Cafiso, D.S., 2006. Recent advances and applications of site-directed spin labeling. Curr. Opin. Struct. Biol. 16, 644–653. Frazier, A.A., Wisner, M.A., Malmberg, N.J., Victor, K.G., Fanucci, G.E., 2002. Membrane orientation and position of the C2 domain from cPLA2 by sitedirected spin labeling. Biochemistry 41, 6282–6292. Frimer, A.A., Strul, G., Hameiri-Buch, J., Gottlieb, H.E., 1996. Can superoxide organic chemistry be observed within the liposomal bilayer? Free Radic. Biol. Med. 20, 843–852. Fukuzawa, K., 2008. Dynamics of lipid peroxidation and antioxidion of alphatocopherol in membranes. J. Nutr. Sci. Vitaminol. 54, 273–285. Gamliel, A., 2005. Spin-Trapping with New Lipophilic Spin-Traps, Ph.D. Thesis. BarIlan: Ramat Gan, Israel. Gamliel, A., Afri, M., Frimer, A.A., 2008. Determining radical penetration of lipid bilayers with new lipophilic spin traps. Free Radic. Biol. Med. 44, 1394–1405. Gregoriadis, G., 1984. Liposome Technology, Vol. I–III. CRC Press, Boca Raton. Herenyi, L., Veres, D., Békási, S., Voszka, I., Módos, K., Csík, G., Kaposi, A.D., Fidy, J., 2009. Location of mesoporphyrin in liposomes determined by site-selective fluorescence spectroscopy. J. Phys. Chem. B 113, 7716–7724. Holmes, D.L., Lightner, D.A., 1996. Synthesis and acidity constants of 13CO2Hlabelled dicarboxylic acids. pKas from 13C NMR. Tetrahedron 52, 5319–5338. Hong, M., 2006. Solid-state NMR studies of the structure, dynamics, and assembly of b-sheet membrane peptides and a-helical membrane proteins with antibiotic activities. Acc. Chem. Res. 39, 176–183. Hubbell, W.L., Altenbach, C., 1994. Investigation of structure and dynamics in membrane proteins using site directed spin labeling. Curr. Opin. Struct. Biol. 4, 566–573. Huster, D., Muller, P., Arnold, K., Herrmann, A., 2001. Dynamics of membrane penetration of the fluorescent 7-nitrobenz-2-oxa-1,3-diazol-4-yl (NBD) group attached to an acyl chain of phosphatidylcholine. Biophys. J. 80, 822–831. Jämbeck, J.P.M., Lyubartsev, A.P., 2012. Derivation and systematic validation of a refined all-atom force field for phosphatidylcholine lipids. J. Phys. Chem. B 116, 3164–3179. Khurana, J.M., Sahoo, P.K., Maikap, G.C., 1990. An international journal for rapid communication of synthetic organic chemistry. Syn. Commun. 20, 2267–2271. Komljenovic, I., Marquardt, D., Harroun, T.A., Sternin, E., 2010. Location of chlorhexidine in DMPC model membranes: a neutron diffraction study. Chem. Phys. Lipids 163, 480–487. Ku9 cerka, N., Kisekev, M.A., Balgavy, P., 2004. Determination of bilayer thickness and lipid surface area in unilamellar dimyristoylphosphatidylcholine vesicles from small-angle neutron scattering curves: a comparison of evaluation methods. Eur. Biophys. J. 33, 328–334.

118

M. Afri et al. / Chemistry and Physics of Lipids 184 (2014) 105–118

Ku9 cerka, N., Liu Chu, Y.N., Petrache, H.I., Tristram-Nagle, S., Nagle, J.F., 2005. Structure of fully hydrated fluid phase DMPC and DLPC lipid bilayers using X-ray scattering from oriented multilamellar arrays and from unilamellar vesicles. Biophys. J. 88, 2626–2637. Ku9 cerka, N., Marquardt, D., Harroun, T.A., Nieh, M.-P., Wassall, S.R., de Jong, D.H., Schafer, L.V., Marrink, S., Katsaras, J., 2010. Cholesterol in bilayers with PUFA chains: doping with DMPC or POPC results in sterol reorientation and membrane-domain formation. J. Biochem. 49, 7485–7493. Ku9 cerka, N., Nieh, M.-P., Katsaras, J., 2011. Fluid phase lipid areas and bilayer thicknesses of commonly used phosphatidylcholines as a function of temperature. Biochim. Biophys. Acta 2761–2771. Lodyato, V.I., Yurkova, I.L., Sorokin, V.L., Shadyro, O.I., Dolgopalets, V.I., Kisel, M.A., 2004. Novel (3,5-di-tert-butyl-2-hydroxy-phenylcarbamoyl)-alkanoic acids as potent antioxidants. Bioorg. Med. Chem. Lett. 14, 4253–4256. March, J., 1992. Advanced Organic Chemistry: Reactions, Mechanisms, and Structures, fourth ed. Wiley, New York. Marquardt, D., Williams, J.A., Kinnun, J.J., Ku9 cerka, N., Atkinson, J., Wassall, S.R., Katsaras, J., Harroun, T.A., 2014. Dimyristoyl phosphatidylcholine: a remarkable exception to a-tocopherol’s membrane presence. J. Am. Chem. Soc. 136, 203–210. Menger, F.M., Keiper, J.S., Caran, K.L., 2002. Depth-profiling with giant vesicle membranes. J. Am. Chem. Soc. 124, 11842–11843. Menger, F.M., Richardson, S.N., Wood Jr., M.G., Sherrod, M.J., 1989. Chain-substituted lipids in monomolecular films. Effect of polar substituents on molecular packing. Langmuir 5, 833–838. Moreau, J.J.E., Vellutini, L., Man, M.W.C., Bied, C., Dieudonne, P., Bantignies, J.-L., Sauvajol, J.L., 2005. Lamellar bridged silsesquioxanes: self-assembly through a combination of hydrogen bonding and hydrophobic interactions. Chem. Eur. J. 11, 1527–1537. Mourelatou, E.A., Libster, D., Nir, I., Hatziantoniou, S., Aserin, A., Garti, N., Demetzos, C., 2011. Type and location of interaction between hyperbranched polymers and liposomes. Relevance to design of a potentially advanced drug delivery nanosystem (aDDnS). J. Phys. Chem. B 115, 3400–3408. Nielsen, R.D., Che, K., Gelb, M.H., Robinson, B.H., 2005. A ruler for determining the position of proteins in membranes. J. Am. Chem. Soc. 127, 6430–6442. Oda, H., Kobayashi, T., Kosugi, M., Migita, T., 1995. Synthesis of 5,12-dihete derivative by palladium-catalyzed ternary coupling between vinylic halide, a vinylic tin compound, and norbornadiene. Tetrahedron 51, 695–702. Papahadjopoulos, D., 1978. Liposomes and Their Uses in Biology and Medicine. Academy of Sciences, New York. Postupalenko, V.Y., Shvadchak, V.V., Duportail, G., Pivovarenko, V.G., Klymchenko, A. S., Mély, Y., 2011. Monitoring membrane binding and insertion of peptides by two-color fluorescent label. Biochim. Biophys. Acta 1808, 424–432.

Prosser, R.S., Evanics, F., Kitevski, J.L., Patel, S., 2007. The measurement of immersion depth and topology of membrane proteins by solution state NMR. Biochim. Biophys. Acta 1768, 3044–3051. Rao, A.V.R., Mysorekar, S.V., Yadav, J.S., 1987. Synthesis of 1,3-diols: its application to the synthesis of a-lipoic acid. Synth. Commun. 17, 1339–1347. Reichardt, C., 1965. Empirical parameters of the polarity of solvents. Angew. Chem. Int. Ed. 4, 29–40. Reichardt, C., 1994. Solvatochromic dyes as solvent polarity indicators. Chem. Rev. 94, 2319–2358. Reichardt, C., Welton, T., 2011. Solvents and Solvent Effects in Organic Chemistry, fourth ed. Wiley-VCH Weinhein, Germany Chapter 6 and 7. Ruíz-Gutiérrez, V., Prada, J., Pérez-Jiménez, F., 1993. Determination of fatty acid and triacylglycerol composition of human very-low-density lipoproteins. J. Chromatogr. B 622, 117–124. Sassaroli, M., Vauhkonen, M., Perry, D., Eisinger, J., 1990. Lateral diffusivity of lipid analogue excimeric probes in dimyristoylphosphatidylcholine bilayers. Biophys. J. 57, 281–290. Serbinova, E., Kagan, V., Han, D., Packer, L., 1991. Free radical recycling and intramembrane mobility in the antioxidant properties of alpha-tocopherol and alpha-tocotrienol. Free Radic. Biol. Med. 10, 263–275. Shachan-Tov, S., Afri, M., Frimer, A.A., 2010. A reinvestigation of the reaction of coumarins with superoxide in the liposomal bilayer correlation between depth and reactivity. Free Radic. Biol. Med. 49, 1516–1521. Takahashi, M., Tsuchiya, J., Niki, E., 1989. Scavenging of radicals by vitamin E in the membranes as studied by spin labeling. J. Am. Chem. Soc. 111, 6350–6353. Tanaka, A., 1959. Anodic synthesis of fatty acids IV. Synthesis of chemical constituents of Conifer wax. Yakugaku Zasshi 79, 1327–1331 cited by Chem. Abstr. 1960, 54, 4381. Tulloch, A.P., 1977. Deuterium isotope effects and assignment of 13C chemical shifts in spectra of methyl octadecanoate and the sixteen isomeric oxooctadecanoates. Can. J. Chem. 55, 1135–1142. Voinov, M.A., Kirilyuk, I.A., Smirnov, A.I., 2009. Spin-labeled pH-sensitive phospholipids for interfacial pKa determination: synthesis and characterization in aqueous and micellar solutions. J. Phys. Chem. B 113, 3453–3460. Wassall, S., Wang, L., McCabe, R., Ehringer, W., Stillwell, W., 1991. Electron spin resonance study of the interaction of alpha-tocopherol with phospholipid model membranes. Chem. Phys. Lipids 60, 29–37. Weitman, H., Roslaniec, M., Frimer, A.A., Afri, M., Freeman, D., Mazur, Y., Ehrenberg, B., 2001. Solvatochromic effects in the electronic absorption and nuclear magnetic resonance spectra of hypericin in organic solvents and in lipid bilayers. Photochem. Photobiol. 73, 110–118.