A General Method for Quantification and Discovery of Acyl Groups Attached to Acyl Carrier Proteins in Fatty Acid Metabolism Using LC-MS/MS

Acyl-ACPs that enable fatty acid biosynthesis were uniquely profiled and quantified, resulting in discovery of novel ACP products with unknown function and providing a method for rigorous comparisons.

Abstract

Acyl carrier proteins (ACPs) are the scaffolds for fatty acid biosynthesis in living systems, rendering them essential to a comprehensive understanding of lipid metabolism. However, accurate quantitative methods to assess individual acyl-ACPs do not exist. We developed a robust method to quantify acyl-ACPs to the picogram level. We successfully identified acyl-ACP elongation intermediates (3-hydroxyacyl-ACPs and 2,3-trans-enoyl-ACPs) and unexpected medium-chain (C10:1, C14:1) and polyunsaturated long-chain (C16:3) acyl-ACPs, indicating both the sensitivity of the method and how current descriptions of lipid metabolism and ACP function are incomplete. Such ACPs are likely important to medium-chain lipid production for fuels and highlight poorly understood lipid remodeling events in the chloroplast. The approach is broadly applicable to type II fatty acid synthase systems found in plants and bacteria as well as mitochondria from mammals and fungi because it capitalizes on a highly conserved Asp-Ser-Leu-Asp amino acid sequence in ACPs to which acyl groups attach. Our method allows for sensitive quantification using liquid chromatography–tandem mass spectrometry with de novo–generated standards and an isotopic dilution strategy and will fill a gap in our understanding, providing insights through quantitative exploration of fatty acid biosynthesis processes for optimal biofuels, renewable feedstocks, and medical studies in health and disease.

INTRODUCTION

The synthesis of fatty acyl chains is essential for the production of storage and membrane lipids as well as functional molecules that modulate gene expression or contribute to protein activity. Studies on acyl chains span many themes, from disease and nutrition to the demand for renewable fuels and feedstocks (Uauy et al., 2000; Peralta-Yahya et al., 2012; Zhu et al., 2016; Vanhercke et al., 2019). Carbon chains of up to 18 carbons in length are produced during fatty acid biosynthesis (Li-Beisson et al., 2013) on an acyl carrier protein scaffold (Overath and Stumpf, 1964) that is connected to the acyl chain through linkage of a 4′-phosphopantetheine group to a Ser residue found in the acyl carrier protein (ACP; Figure 1A). The acyl chains are elongated by a series of four enzymatic steps that link ACP intermediates and form a cycle that results in the addition of two carbons and four hydrogens. Two distinct types of fatty acid synthase (FAS) complexes perform the repeated cycle of ketoacyl synthesis, reduction, dehydration, and a second reduction producing long-chain saturated hydrocarbons (Figure 1B). Yeast and mammals use a single, large multifunctional FAS protein (type I) that includes multiple domains for the enzymatic steps and an ACP region (Bressler and Wakil, 1961; Hsu et al., 1965), whereas fatty acid synthesis in plants and bacteria occur through the concerted action of individual proteins within a type II FAS complex and a distinct ACP that is 9 to 15 kD in size (Alberts et al., 1963; White et al., 2005; Cronan, 2014).

Figure 1.

Acyl-ACP Structure and Function.

(A) Acyl-ACP is a 9- to 15-kD protein with a 4′-phosphopantetheine arm attaching a fatty acyl chain to the protein backbone. The structure of acyl-ACP contains a highly conserved sequence of amino acids flanking the Ser that attaches to phosphopantetheine. Peptide hydrolysis at aspartate residues produces an acyl molecular species that can be processed for characteristic fragments by mass spectrometry that have precursor-product losses of 315.1 and 413.1 D.

(B) ACPs are central to fatty acid synthesis with the process of fatty acid biosynthetic elongation of an acyl chain by two carbons taking place on ACP intermediates through a series of enzymatic steps that are repeated and conclude with acyl transfer to glycerol-3-phosphate or lyso-phosphatidic acid for production of chloroplast lipids or hydrolysis by a thioesterase. Nonesterified fatty acids, produced through hydrolysis, are generally exported from the chloroplast, esterified to CoA, and used for lipid biosynthesis in the endoplasmic reticulum (ER).

In plants, the production of acyl chains takes place predominantly in the chloroplast, and to a lesser extent in mitochondria, using acyl-ACP intermediates in both instances (Ohlrogge et al., 1979; Chuman and Brody, 1989). The chain elongation process is terminated in the chloroplast by acyl-ACP–dependent acyltransferases or by thioesterase reactions; the former contribute acyl groups for lipid biosynthesis in the chloroplast, while the latter release a nonesterified fatty acid from the ACP that can be later exported. In both cases, the ACP substrate pool is regenerated for further fatty acid biosynthetic reactions. Nonesterified fatty acids can be activated to acyl-CoA molecules (acyl-CoAs) outside the chloroplast to make glycerolipids for storage oil production in the endoplasmic reticulum, waxes, surface lipids, or membrane biogenesis (Li-Beisson et al., 2013).

Given the central role of acyl-ACPs in fatty acid biosynthesis, their quantitative analysis can help our understanding of lipid metabolism; however, the acyl group is a small percentage of the total acyl-ACP weight (usually much less than 3%), and current methods are confined by biochemical approaches that analyze intact proteins or their subunits. After ACP was demonstrated to be the acyl carrier for fatty acid synthesis (Goldman et al., 1963), numerous studies examined their expression (Hlousek-Radojčić et al., 1992; Bonaventure and Ohlrogge, 2002), subcellular roles (Brody et al., 1997; Wada et al., 1997; Cronan et al., 2005; Witkowski et al., 2007), and the impact of ACP sequence on function and growth (De Lay and Cronan, 2007; Zhu et al., 2019). However, studies quantifying the amount of acyl-ACPs are uncommon.

Levels of acyl-ACPs can be currently measured based on urea polyacrylamide gel separation and immunodetection (Post-Beittenmiller et al., 1991); however, multiple isoforms cannot be resolved (Ohlrogge and Kuo, 1985), and only a subset of all acyl-ACPs are detected by antibodies and gel gradients, limiting the extent of quantification. Focusing more specifically on the acyl component, Kopka et al. (1995) developed an assay for individual acyl-ACP levels with gas chromatography linked to mass spectrometry by generating butylamide derivatives. Levels of spinach (Spinacia oleracea) leaf acyl-ACPs qualitatively matched those inferred from densitometry of urea-based gels. Unfortunately, the chemical derivatization was not specific to acyl-ACP thioester bonds, requiring the meticulous removal of acyl-CoAs and other thioesters—present at greater levels—by anion exchange chromatography prior to derivatization. Moreover, short-chain ACPs cannot be detected through this strategy.

We report a new method to quantitatively analyze acyl-ACPs with tandem mass spectrometry (MS/MS) based on a highly conserved contiguous amino acid region that flanks the attachment of the acyl chain to the ACP. Enzymatic cleavage resulted in an acyl chain connected to a phosphopantetheinyl group and a tripeptide that was sensitively quantified by liquid chromatography (LC)-MS/MS. ¹⁵N-Labeled ACP standards were enzymatically synthesized de novo for absolute quantification of acyl-ACP levels using an isotopic dilution strategy. The method identified 3-hydroxyacyl-ACP and 2,3-trans-enoyl-ACP, two of the three acyl-ACP elongation intermediates for each two-carbon acyl-ACP product of the fatty acid biosynthetic cycle. We also detected unanticipated medium-chain unsaturated acyl-ACPs that to date have no known function in plant metabolism. Tests with false flax (Camelina sativa, hereafter Camelina), an emerging oilseed crop, found ∼26 pmol/mg fresh weight (FW) of acyl-ACPs in developing seeds, present mostly in the terminal ACP product 18:1-ACP. By contrast, leaves had much lower levels (∼3 pmol/mg FW). This report represents a comprehensive approach to profile acyl-ACP products with the absolute quantification of resolved individual acyl-ACP levels. The application of this method to profile acyl-ACP products is a critical step to improve our understanding of fatty acid metabolism and lipid regulation.

RESULTS

Acyl-ACP Standards Were Generated for Absolute Quantitation

Because acyl-ACP standards are not commercially available, isotope-labeled standards were generated using a 4′-phosphopantetheinyl transferase from Bacillus subtilis, Sfp transferase (Sfp; Figure 2), overexpressed in Escherichia coli. The apo-ACP substrate was also overexpressed in E. coli grown in Luria-Bertani (LB) medium or in a minimal medium supplemented with ¹⁵NH₄Cl to produce unlabeled and ¹⁵N-apo-ACP, respectively. The Sfp and ACP products were purified by nickel affinity chromatography. Gel densitometry estimated the range of apo-ACP purity (79% ± 5% [sd], n = 36); the remaining 21% was phosphopantetheinylated. Protein concentration was calculated from the absorbance and molar extinction coefficients. Unlabeled- and ¹⁵N-apo-ACPs (100 to 250 µM) and acyl-CoAs (300 to 500 µM) served as substrates that were enzymatically reacted with the Sfp (25 µM) to produce individual acyl-ACPs.

Figure 2.

Enzymatic Synthesis of Acyl-ACP Standards Using Sfp Transferase.

Sfp covalently transfers the acyl-4′-phosphopantetheine prosthetic group (highlighted in blue) from acyl-CoA to conserved Ser sites on apo-ACP. Nonesterified (holo), saturated acyl chains with lengths of 2 to 18 carbons, and 16:1 and 18:1 acyl-ACP standards were synthesized.

Acyl-CoA substrates with varying chain lengths were used to test the enzymatic reaction efficiency with various buffer conditions and concentrations of DTT, MgCl₂, and MnCl₂. Buffering to pH 6.5 with 50 mM MOPS improved product formation, in agreement with prior research on phosphopantetheinyl transferases (Quadri et al., 1998; Sánchez et al., 2001; Joshi et al., 2003). Inclusion of DTT (1 to 5 mM) improved reaction efficiency in some instances but caused product degradation in others. We therefore substituted DTT for the more suitable tris(2-carboxyethyl)phosphine (TCEP), provided at 4 mM, to improve stability over a greater pH range and enhance disulfide selectivity. Additionally, we added 10 mM MgCl₂ and MnCl₂ to achieve near-quantitative conversion of apo-ACP to short-chain (C2) and medium-chain (C8) acyl-ACP within a 3-h reaction time at 37°C and increase long-chain (C18) reaction conversion, although not to completion (Supplemental Figure 1). The addition of MgCl₂ and MnCl₂ resulted in partial precipitation of medium- and long-chain acyl-CoAs (i.e., C10 to C18-CoA; Constantinides and Steim, 1986) that was alleviated by DMSO (20% [v/v]) and Tween 20 (1% [w/v]) without impact on enzyme proficiency (Supplemental Figure 2). Using these conditions (Supplemental Table 1), we consistently achieved yields greater than 90% for all acyl chains, except for malonyl-ACP as it could not be reliably obtained due to the easily decarboxylated and thus unstable malonyl-CoA substrate.

ACPs from Many Organisms Share a Conserved Sequence That Is Efficiently Cleaved with the Endoproteinase Asp-N

Elongating acyl chains are the component of interest within fatty acid synthesis; however, the acyl group is small in mass relative to the protein portion of acyl-ACPs. We hypothesized that methods to remove much of the protein chain would benefit downstream quantitative analyses of the acyl composition. Although the amino acid sequences in ACPs are diverse and vary between isoforms and across species, a highly conserved region fortuitously flanks the Ser residue that joins the protein to the phosphopantetheine group. Vascular plants, algae, many bacteria, and mitochondrial ACPs from yeast, insects, and mammals possess a four–amino acid sequence (Asp-Ser-Leu-Asp; DSLD) surrounding the acyl attachment site (Figure 1A; Table 1). Therefore, proteolytic hydrolysis adjacent to aspartate residues with the protease Asp-N should produce a three–amino acid peptide linked to the 4′-phosphopantetheine and acyl group (Figure 1A).

Table 1. Highly Conserved Amino Acid Sequence in ACPs.

Description of Species/Isoform	Locus/Protein Name	Common Name	Conserved Sequence	Ser Residue
Apis mellifera	XP_016766950	Honeybee	LGLDSLDHVE	144
Arabidopsis thaliana ACP1	At3g05020	Arabidopsis	LGADSLDTVE	93
Arabidopsis thaliana ACP2	At1g54580	Arabidopsis	LGADSLDTVE	92
Arabidopsis thaliana ACP3	At1g54630	Arabidopsis	LGADSLDTVE	91
Arabidopsis thaliana ACP4	At4g25050	Arabidopsis	LGADSLDTVE	88
Arabidopsis thaliana ACP5	At5g27200	Arabidopsis	LGADSLDTVE	94
Arabidopsis thaliana mtACP1	At2g44620	Arabidopsis	LGLDSLDTVE	79
Arabidopsis thaliana mtACP2	At1g65290	Arabidopsis	LGLDSLDTVE	83
Bacillus subtilis (168)	NP_389474	Bacillus	LGADSLDVVE	37
Brassica rapa	CAA49802	Brassica	LGADSLDTVE	90
Camelina sativa ACP1	XP_010464003	Camelina	LGADSLDTVE	90
Camelina sativa ACP2a	XP_010414911	Camelina	LGADSLDTVE	92
	XP_010470364	Camelina	LGADSLDTVE	95
Camelina sativa ACP3a	XP_010414908	Camelina	LGADSLDTVE	90
	XP_010511204	Camelina	LGADSLDTVE	89
Camelina sativa ACP4	XP_010438929	Camelina	LGADSLDTVE	89
Camelina sativa ACP5	XP_010455149	Camelina	LGADSLDTVE	96
Camelina sativa mtACP1	XP_010518065	Camelina	LGLDSLDTVE	79
Camelina sativa mtACP2	XP_010470432	Camelina	LGLDSLDTVE	83
Camelina sativa mtACP3	XP_010441433	Camelina	LSLDSLDKVE	86
Chlamydomonas reinhardtii - ACP2	XP_001693782	Chlamy	LGADSLDTVE	72
Chlamydomonas reinhardtii - mtACP1	XP_001699275	Chlamy	LGLDSLDVVE	85
Cuphea lanceolata	P52411	Cuphea	LGADSLDTVE	95
Danio rerio	NP_001292499	Zebrafish	LGLDSLDQVE	109
Enterobacterales	WP_015833846	Bacteria	LGADSLDTVE	37
Escherichia coli IAI39	YP_002408032	E. coli	LGADSLDTVE	37
Escherichia coli (strain K12)	NP_415612	E. coli	LGADSLDTVE	37
Escherichia coli (O83:H1)	YP_006119417	E. coli	LGADSLDTVE	37
Escherichia coli (O157:H7)	BAB34895	E. coli	LGADSLDTVE	47
Escherichia coli (UMN026)	CAR12479	E. coli	LGADSLDTVE	37
Galleria mellonella	XP_026761684	Wax moth	LGLDSLDHVE	108
Gallus gallus	XP_004945320	Chicken	LGLDSLDQVE	106
Glycine max	XP_003536101	Soybean	LGADSLDTVE	99
Helianthus annuus	ADV16365	Sunflower	LGADSLDTVE	94
Homo sapiens	NP_004994	Human	LGLDSLDQVE	112
Homo sapiens	XP_011544158	Human	LGLDSLDQVE	143
Mus musculus	NP_082453	Mouse	LGLDSLDQVE	112
Proteobacteria	WP_003857954	Bacteria	LGADSLDTVE	37
Ricinus communis	XP_002512285	Castor	LGADSLDTVE	91
Saccharomyces cerevisiae (strain S288C)	CAA82036	Yeast	LGLDSLDTVE	82
Shewanella oneidensis (MR-1)	NP_718356	Shewanella	LGADSLDTVE	37
Synechocystis (multiple)	WP_010871942	Synechocystis	LGADSLDTVE	38
Spinacia oleracea	XP_021844779	Spinach	LGADSLDTVE	94
Zea mays	ACG49250	Zea	IGADSLDTVE	94

ACPs associated with type II FASs frequently contain a contiguous region of four amino acids (DSLD, underlined). The Ser residue provides the point of attachment to the acyl limb described in Figure 1A. For most plant ACPs, only one chloroplast isoform is shown except where noted. Bacteria including Bacillus subtilis, Shewanella oneidensis, Synechocystis common sequence, E. coli (K12 strains), Enterobacterales, and Proteobacterial sequences were inspected. Yeast, animal, and other mitochondrial examples of ACP conserved sequence are also presented. Sequences for ACP protein alignment were obtained from National Center for Biotechnology Information protein database and are listed alphabetically by species.

^aMultiple accessions provided differ in chromosomal location and sequence yet have been putatively annotated as the same ACP.

We optimized ACP digestion using visual detection from the chromophore BODIPY-FL-N-(2-aminoethyl)maleimide-S-ACP (BODIPY-ACP) conjugate as described in the Methods with several commercially available forms of the Asp-N protease. Digestion of synthesized acyl-ACP standards was most efficient when the standards were purified away from other reaction components by trichloroacetic acid (TCA) precipitation, possibly due to substrate inhibition of Asp-N proteolytic activity. Recovery of the acyl-ACP standards postsynthesis was assessed by gel densitometry (69% ± 14% [sd], n = 9; Supplemental Figure 3). Acyl-ACP standards were digested with Asp-N protease at enzyme-to-acyl-ACP ratios of 1:20, 1:50, and 1:100 at various pHs, temperatures, and times. The reaction products were analyzed by SDS-PAGE to establish an adequate reaction system. Although short-chain acyl-ACPs could be easily digested in less time and with less enzyme, obtaining complete digestion of medium-to-long chains required a 1:20 ratio of enzyme to substrate for 16 h at 37°C (Supplemental Figure 4). All digestions were quenched by the addition of methanol (50% of final concentration).

Acyl-ACP–Digested Products Can Be Sensitively Detected by LC-MS/MS

Digested acyl-ACP standards and biological samples were separated by reversed phase chromatography on a C18 column and analyzed by mass spectrometry in positive mode (i.e., z = 1, m/z + 1 adduct), using a liquid chromatography–tandem mass spectrometer as described in the Methods (Figure 3A). We used the masses calculated from chemical composition along with the multiple reaction monitoring (MRM) mode of analysis of the MS/MS to confirm the products of acyl-ACP fragmentation and optimize instrument performance. We consistently observed two mass losses for all acyl-ACP digestion products, indicating that they were not specific to the changing size in acyl chain but instead to components of the redundant 4′-phosphopantetheine and three–amino acid sequence (Figure 1A). A product ion that differed from the precursor by 315.1 D indicated the loss of the three amino acids (DSL). The additional loss of phosphate brought the difference in mass to 413.1 D and resulted in the primary product ion observed by mass spectrometry (Figure 3B). Malonyl-ACP had an additional product ion with a difference in mass of 457.1 D, which indicated the loss of the DSL, phosphate, and the carboxyl from the malonyl group corresponding to the main product ion generated for this molecule. The ¹⁵N-labeled standards are 3 D heavier, giving a precursor-product ion difference of 318.1 and 460.1 D. We used the changes in abundance to optimize the declustering potential (DP), collision energy (CE), and collision cell exit potential from standards and acyl-ACPs isolated from plant biomass (Figure 3C). We observed spectra with well-defined peaks that were evenly spaced to represent the change in acyl mass by successive increments of C₂H₄, consistent with optimal chromatography and detection on the mass spectrometer of acyl-ACP–digested products. We identified all acyl species, from acetyl-ACP to 18 carbon acyl chains (i.e., 18:0-ACP), in addition to nonesterified (i.e., holo) and apo-ACP. We established the dynamic range of our method (up to three orders of magnitude) through comparison of integrated peak areas for each acyl-ACP, with the exception of holo- and malonyl-ACP due to dimerization and instability, from standards and Camelina samples: as low as femtomole quantities (i.e., picograms of protein) could be quantified accurately (Figure 3C; Supplemental Figure 5). Since pooled acyl-ACPs are found at low concentrations (10 ng/mg FW in spinach chloroplast; Ohlrogge et al., 1979), saturation of the detector is unlikely to become an issue for most preparations.

Figure 3.

Mass Spectral Detection and Analysis of Acyl-ACPs.

(A) Chromatographic properties of acyl-ACP standards. Asp-N proteolytically digested partial acyl-ACPs are separated according to the length and properties of the acyl chain on a reversed phase column by LC and detected by triple quadrupole mass spectrometer. Labels above peaks represent the acyl species in the acyl-ACP molecules. cps, counts per second.

(B) MS/MS fragmentation product ion scan of Asp-N proteolytically digested 15:0-ACP standard indicating the major losses. cps, counts per second.

(C) Optimal mass spectrometry parameters for acyl-ACP detection by MRM. Responses are represented as the slope (ratio of peak areas per pmol analyte) of the calibration curve for each ACP species. Linear regression (R²) weighted by 1/Y for all samples. The sd of the lowest detectable levels and slope of the calibration curve were used to establish the limit of detection (LOD = 3 × SD/slope) and quantification (LOQ = 10 × SD/slope). CE, collision energy; CXP, collision cell exit potential; DP, declustering potential; na, not applicable.

Total Acyl-ACP Levels in Camelina Seeds Are Significantly Greater Than in Leaves

Isotope dilution, a general strategy that relies upon internal standard addition for derivation of calibration curves, is the benchmark for quantitation by mass spectrometry. Isotopically labeled internal standards provide chemically equivalent ions distinguished from sample metabolites, or analytes, by a change in mass corresponding to the isotope label(s). Our method shifts the mass by ∼3 D due to the substitution of three ¹⁵N atoms in place of ¹⁴N. The addition of internal standards at an early stage during sample preparation allows one to normalize data and rectify sample-to-sample variations in analyte recovery. In addition, the chemical equivalency of isotopically labeled standards can account for differences in observed analyte response factors and address the exposure to differential matrix effects, making isotope dilutions the superior approach. We therefore used an isotope dilution strategy to quantify acyl-ACP levels in Camelina seeds and leaves. Analyte acyl-ACPs from plant biomass were spiked with internal standards before extraction and then quantified. Total ACP amounts detected in seeds were ∼26 pmol/mg FW and were composed predominantly of 18:1-ACP (Figure 4). Camelina leaves exhibited approximately ninefold lower levels overall than seeds (i.e., ∼3 pmol/mg FW), consistent with calculated estimates from the literature (∼1 to 2 pmol/mg FW calculated from spinach leaves; Ohlrogge et al., 1979) and of comparable acyl composition (Post-Beittenmiller et al., 1991).

Figure 4.

Acyl-ACP Levels in Camelina Seeds and Leaves.

Acyl-ACPs from green seeds 61 DAS (n = 11 to 15; except 16:1, 18:0, and 18:1, n = 4) and leaves (28 DAS; n = 16 to 19; except 16:1, n = 4; 18:0, n = 6; and 18:1, n = 12) were quantified using internal standards. Lower replication of saturated and unsaturated long chains reflect initial digestion inefficiencies in sample preparation. The means and sds are shown for each acyl species.

Two Acyl-ACP Intermediates of the Fatty Acid Biosynthetic Cycle Were Identified and Previously Undescribed Unsaturated Acyl-ACPs Were Discovered

The process of acyl chain extension in fatty acid synthesis is completed by a cycle that generates three additional acyl-ACP elongation intermediates reflecting the ketoacyl condensing, reducing, and dehydrating steps before the final reduced acyl-ACP. Although these intermediates are believed to be present at low quantities, we asked whether our method could detect them. Since the intermediates are themselves ACP products that differ only in the acyl structure and composition, the developed methods were presumed applicable, although potentially limited by the instrument sensitivity.

Analysis of seeds detected several additional acyl-ACP peaks. We calculated the molecular weights for the elongation intermediates assuming a loss of 413.1 D (like other acyl-ACPs measured). We identified two acyl-ACP elongation intermediates for each acyl chain-length form: 3-hydroxyacyl-ACPs and 2,3-trans-enoyl-ACPs. The MRM list (Supplemental Table 2) also included the exact masses for 3-ketoacyl-ACPs, the first intermediate in the fatty acid biosynthetic cycle (Figure 1B), even though this intermediate was not detected. Developing seeds accumulated 3-hydroxyacyl-ACPs to similar amounts as most reduced acyl-ACPs (Figure 5A) with higher levels of C14-hydroxyacyl-ACP that may be involved in lipid A–like molecule production in the mitochondria (Wada et al., 1997; Li et al., 2011). Peaks for the 2,3-trans-enoyl-ACP were lower in area, generally less than 2.0% of total (C18-enoyl; Figures 5B).

Figure 5.

Acyl-ACP Elongation Intermediates Detected from Camelina Developing Seeds.

Results from LC-MS/MS analysis.

(A) Relative peak areas, expressed as percentage, were calculated from all the acyl-ACP species detected. Means and sds are shown based on four independent sample preparations.

(B) Chromatographic detection of unanticipated unsaturated acyl-ACPs. The characteristic double peak pattern for the 2,3-trans-enoyl- and desaturated acyl-ACP isomers were observed for two medium-chain acyl-ACPs (C10, dark blue; C14, green). Polyunsaturated C16-ACP (16:3-ACP, black) was also detected. cps, counts per second.

2,3-trans-Enoyl-ACP and equivalent chain-length single desaturated acyl-ACPs are isomeric. Therefore, isomers such as 16:1-ACP (cis-double bond) and C16-enoyl-ACP (2,3-trans-double bond) cannot be discriminated by mass. Consequently, acyl chain MRM transitions indicated a second peak for 2,3-trans-enoyl acyl-ACP at a different retention time. To conclusively identify the isomers, we digested and processed a standard of 18:1-ACP, resulting in a peak that eluted at the same retention time as one of the two peaks in question (Supplemental Figure 6). This confirmed the designation of each peak, with 2,3-trans-enoyl acyl-ACP characteristically eluting after the cis-unsaturated acyl-ACP as a much smaller peak. 3-Hydroxyacyl-ACP products that differ in mass eluted ahead of the corresponding acyl-ACP as presented in a summary of the retention times in Supplemental Table 3.

Interestingly, owing to the sensitivity of the method, we observed peaks corresponding to several medium-chain unsaturated acyl-ACPs (i.e., C10:1 and C14:1) and a significant amount of polyunsaturated acyl-ACP (16:3) in seeds and leaves (Figure 5B) that had not been previously described. The paired MS/MS transitions, retention times, and peak shapes all supported the identification of these species. The retention time offset (Figure 5B) was similar to measured differences for 9-cis-18:1-ACP (Supplemental Figure 6) and 7-cis-16:1-ACP and suggested that these unsaturated forms likely contain cis-double bonds. Enoyl-ACPs contain trans-double bonds and have a predicted near-linear, three-dimensional structure and therefore more closely approximate saturated ACPs with respect to their physical properties. cis-Unsaturated-acyl groups are commonly generated within living systems, although they are not believed to be biosynthetic intermediates on FAS assembly lines; thus, the roles in metabolism for these molecules remain to be elucidated.

DISCUSSION

Development of an Acyl-ACP Profiling Method Based on a Conserved Contiguous Peptidyl Region

Acyl chains represent 90 to 95% of the carbon in glycerolipids; thus, the production and movement of fatty acids is central to lipid metabolism. We developed a sensitive method to identify and quantify acyl-ACPs that can complement analysis of acyl-CoAs to provide a more complete picture of the connectivity between fatty acid biosynthesis and lipid assembly (Allen, 2016a). The growing and nascent acyl chains are tethered to the ACP via a 4′-phosphopantetheinyl linker and to a Ser residue that is part of a conserved four–amino acid phosphopantetheine attachment site (Asp-Ser-Leu-Asp; DSLD). Acyl-ACPs are present in cells in meager quantities and are made from different ACP isoforms. Each can be reduced with an aspartyl protease to three–amino acid peptides linked to 4′-phosphopantetheine that differ only in the attached acyl chains. This effectively eliminates protein size and sequence as a source of variation that otherwise convolute the population of a single acyl-chain species across multiple possible carriers and enables their detection in forms that are convenient for mass spectrometric analysis.

In comparison to previous published attempts, our digestion and LC-MS/MS–based analysis does not require antibodies, multiple urea-PAGE gels, or derivatization techniques and requires only milligram levels of biological material. Acyl-ACPs that are present at picogram/milligram FW quantities can be readily detected without interference from acyl chains esterified to lipids or CoA species that are more abundant in biomass. In addition, the use of the phosphopantetheinyl transferase Sfp, from B. subtilis, which tolerates a wide variety of substituents added onto its canonical substrate, 4′-phosphopantetheine, offered us a one-step synthesis of acyl-ACPs starting with acyl-CoAs and ACP (Figure 2; Lambalot et al., 1996). The approach circumvents a more typical, two-step enzymatic synthesis beginning with production of holo-ACP, often with the E. coli phosphopantetheinyl transferase AcpS, followed by acylation with acyl-ACP synthetase to generate acyl-ACPs (Zornetzer et al., 2006).

Acyl-ACP Profiling Reveals Unanticipated Acyl-ACP Groups with Implications for Lipid Metabolism in Oilseeds

We quantified the absolute levels of individual acyl-ACPs containing up to 18 carbons. 18:1-ACP was present in the greatest quantities in seeds and may indicate that fatty acid biosynthesis is not a bottleneck for lipid production in oilseeds but could be important to regulation (Andre et al., 2012). In addition, we identified two intermediates of the fatty acid biosynthetic cycle: 3-hydroxyacyl-ACP and 2,3-trans-enoyl-ACP. Interestingly, our approach elucidated other unsaturated acyl-ACPs in addition to 18:1, including medium-chain (C10:1 and C14:1) acyl-ACPs with unknown function that had not been reported previously. On a speculative basis, it would be interesting to determine whether plants or other species that exhibit enhanced medium-acyl chain lipid production also accumulate higher levels of these ACP forms. This might indicate differences in the fatty acid biosynthetic machinery and offer potential targets for metabolic engineering. We also observed higher degrees of unsaturation in C16 acyl chains (i.e., 16:3-ACP; Figure 5), which may be relevant to lipid remodeling in the chloroplast. The detection of unanticipated acyl-ACPs indicates that the method can serve as a sensitive and quantitative tool for discovery aspects of fatty acid metabolism.

Absolute Levels of Acyl-ACPs Are Greater in Seeds than Leaves of Oilseeds

The levels of acyl-ACPs in seeds were greater than in leaves. Oilseeds actively produce storage lipids that can represent the largest fraction of seed biomass, whereas leaves make a small amount of lipids primarily for use in membranes (Fan et al., 2014). In agreement with more qualitative studies on spinach leaves (Post-Beittenmiller et al., 1991), our method quantified high levels of long-chain acyl-ACPs in seeds (Figure 4A), predominantly in the form 18:1. This method also separated apo- and holo-ACP pools in Camelina seeds and leaves that were not teased apart in the spinach studies, although the holo-ACP pool could not be accurately quantified. 18:1-ACP is a terminal product of fatty acid de novo synthesis and may indicate that acyl chain production is not a limiting factor in lipid biosynthesis. The presence of an apo-ACP pool may also suggest that ACP abundance does not limit fatty acid production, but perhaps the activation to holo-form and the source of acetyl groups both impact the regulated rate of the fatty acid biosynthetic cycle.

The Role of Acetyl-ACP in Plant Lipid Metabolism and Regulation

In spinach leaves and seeds, 5 to 10% of the total ACP is in an acetylated form (Post-Beittenmiller et al., 1991), which is comparable to the numbers reported here for Camelina tissues (4 and 11% for seed and leaf, respectively; Figure 4). In vitro studies have previously indicated that acetyl-ACP is inefficiently but preferentially used as substrate for condensation reactions catalyzed by the 3-ketoacyl-ACP synthase I isoform rather than 3-ketoacyl-ACP synthase III (Jaworski et al., 1993). It can also be converted to acetyl-CoA by the reversible acetyl-CoA:ACP transacylase reaction. Given the reversibility of transacylase reactions, the steady state levels of acetyl-ACP may reflect demands for acetyl-CoA (Post-Beittenmiller et al., 1991) and regulate the activity of acetyl-CoA carboxylase (Post-Beittenmiller et al., 1991; Andre et al., 2012). In the future, studies on lipid metabolism will benefit from a higher resolution understanding of the fatty acid biosynthetic cycle and should capitalize on the current method with additional transient isotopic labeling studies (Allen, 2016b) to assess synthesis and turnover characteristics.

METHODS

Chemicals, Growth Media, Reagents, and Materials

All chemicals and growth media were purchased from Sigma-Aldrich unless otherwise noted. CoA, malonyl-CoA, acetyl-CoA, butyryl-CoA, hexanoyl-CoA, octanoyl-CoA, decanoyl-CoA, lauroyl-CoA, and myristoyl-CoA were obtained from Sigma-Aldrich. Palmitoyl-CoA, stearoyl-CoA, 16:1(n7)-CoA, and 18:1(n9)-CoA were from Avanti Polar Lipids. BODIPY-FL-N-2-aminoethyl)maleimide (BODIPY), TCEP, and Gelcode stain were from Thermo Fisher Scientific. The 14:0, 15:0, and 16:0-ACPs were gifts from Ed Cahoon (University of Nebraska–Lincoln) at the beginning of the project. SDS-PAGE, agarose gel supplies, and Protein Assay Reagent were from Bio-Rad. DMSO was from Chem-Impex International. The 5-mL HisTrap column and the 5-mL HiTrap desalting columns used for protein purification were from GE Healthcare. Amicon Ultra centrifugal filters used for protein concentration and thin layer chromatography Silica gel 60 plates were from Millipore Sigma. Restriction enzymes, T4 DNA ligase, and DNA polymerase (Finnzymes’ Phusion-Hot Start) were from New England Biolabs. Growth media were from BD Biosciences. Oligonucleotides were purchased from Integrated DNA Technologies and are listed in Supplemental Table 4. PCR cleanup, plasmid miniprep, and gel extraction kits were from Qiagen. Antibiotics were used at the following concentrations: ampicillin or carbenicillin, 100 μg/mL and kanamycin, 50 μg/mL. Camelina (Camelina sativa) plants were grown in the Donald Danforth Plant Center greenhouse. Greenhouse conditions were as follows: temperature, day/night, 22°C/20°C; humidity, 40%; and light/dark, 14 h/10 h. Plants were grown on Berger BM7 35% bark soil (Hummert International) with supplemented fertilizer 3 d per week (Jack’s 15:16:17, JR Peters), concentration 200 ppm nitrogen. Light at 175 μmol photons/m²/s was provided from a combination of 600-W high pressure sodium and 400-W metal halide bulbs.

Microorganisms

Escherichia coli DH5α λpir was a gift for William Metcalf and was used for cloning and plasmid maintenance. E. coli BL21 was purchased from New England Biolabs and was used for protein overexpression. E. coli BL21 (DE3) was purchased from Novagen and used for protein overexpression. E. coli BW25113 was part of the Keio collection (Coli Genetic Stock Center, Yale University). Microorganisms were maintained either on agar selection plates for short-term storage or as a concentrated cell suspension in a 50% medium-glycerol slurry flash-frozen in liquid nitrogen and kept at –80°C for long-term storage. DNA transformations were performed by electroporation of homemade electrocompetent cells.

Plasmids

Plasmid pQE-60-eGFP was constructed by ligation of empty pQE-60 vector (Qiagen) and plasmid pEGFP-CI (Clontech), both of which had been double digested with NheI and XhoI followed by gel purification.

Plasmid pT5Kan (Supplemental Figure 7A) construction began with digestion of pQE-60-eGFP with XhoI and NheI. The gel-purified fragment was ligated to the PCR product of pET-28a (Novagen) amplified by primers pET-28-XhoI-F and pET-28-NheI-R that had been similarly digested and gel purified. The cloned product resulted in an intermediary plasmid that was further modified to remove unnecessary intergenic regions in order to make the vector easier to work with. The truncation was accomplished by PCR amplification of the intermediary plasmid using primers pT5-Kan-NotI-F and pT5-Kan-NotI-R that amplified the lacI-MCS-KanR-ColEI region, leaving only 300 bp separating the lacI and ori-ColEI sequences, as well as provided a NotI site for digestion and ligation of the amplified fragment. The cloned product, named pT5-Kan, was used for cloning and expression of the E. coli acyl carrier protein gene acpP.

Plasmid pT5-Kan-AcpP-His₆ was constructed from the PCR product for acpP amplified from genomic DNA isolated from E. coli MG1655 using primers Acp-His₆-F and Acp-His₆-R and digested with NcoI/BamHI, followed by gel purification and ligation into similarly digested and purified pT5-Kan.

Plasmid pET-Duet-1-Sfp was constructed from NdeI/KpnI-digested PCR product for Sfp amplified from pUC-18 Sfp (a gift from Christopher T. Walsh; Lambalot et al., 1996) using primers Sfp-F and Sfp-R.

Plasmid pET-28a-Sfp-His₆ (Supplemental Figure 7B) was constructed from NcoI/BamHI-digested and gel-purified PCR product for Sfp amplified from pET-Duet-1-Sfp using primers Sfp-His₆-F and Sfp-His₆-R and similarly digested and purified pET-28a.

DNA Sequencing and Analysis

Plasmids were sequenced by Sanger sequencing on an ABI 3730XL capillary sequencer (University of Illinois Biotechnology Center) or ABI 3500 Genetic Analyzer (St. Louis Community College BioBench Contract Research Organization Facility) using BigDye Version 3.0 terminator/enzyme mix and standard protocols. Sequence outputs were assembled and analyzed using the Sequencher 4.6 software (Gene Codes). The primers used in the BigDye reactions were the same as for cloning, except for the primers used to sequence the AcpP-His₆ construct, where we used primers pT5-Kan-F and pT5-Kan-R.

Overexpression and Purification of apo-ACP and Sfp

AcpP from E. coli and Sfp from Bacillus subtilis (Supplemental Table 5) were expressed as C-terminal hexahistidine (His₆) fusion proteins in appropriate E. coli expression strains. For unlabeled protein preparations, the expression strains were grown in LB medium supplemented with kanamycin (50 μg/mL) starting as 1% inoculations from an overnight preculture. Cells were grown at 37°C on an orbital shaker (200 rpm) until mid-log phase (OD₆₀₀ = ∼0.5 to 0.8) before shifting the temperature to 18°C and adding isopropyl-β-d-1-thiogalactopyranoside to a final concentration of 1 mM for overnight induction of expression. Isotopically labeled AcpP was expressed in a modified MOPS minimal medium containing ¹⁵N-labeled ammonium chloride (9.69 mM or 528 mg/L) and kanamycin (50 μg/mL); growth temperature and induction conditions were the same as for the unlabeled preparations. Cells were harvested by centrifugation, resuspended in lysis buffer (10 mM imidazole, 20 mM HEPES, 150 mM NaCl, 1 mM DTT, 10% [v/v] glycerol, 10 mg/mL lysozyme, and protease inhibitor), and lysed by sonication: six cycles of 1 min 50% duty cycle followed by 1 min cooling time at 4°C using a micro-tip probe sonicator. The lysates were then clarified by centrifugation for 20 min at 20,000 rpm before purification. The clarified extracts were applied to a 5-mL HisTrap column (GE Healthcare) using an ÄKTA fast protein liquid chromatography system (GE Healthcare), washed with 20 column volumes of loading/wash buffer (10 mM imidazole, 20 mM HEPES, 150 mM NaCl, 1 mM DTT, and 10% [v/v] glycerol) before elution with elution buffer (250 mM imidazole, 20 mM HEPES, 150 mM NaCl, 1 mM DTT, and 10% [v/v] glycerol). Target protein–containing fractions were concentrated by ultrafiltration using Amicon Ultra Centrifugal Filters (Millipore Sigma) before desalting and buffer exchange into storage buffer (20 mM HEPES, 150 mM NaCl, 1 mM DTT, and 10% [v/v] glycerol) using a 5-mL HiTrap desalting column (GE Healthcare) and a final concentration using the ultrafiltration device described above. Final protein concentrations were determined by measuring absorbance at 280 nm and using molar extinction coefficients calculated using Expasy’s ProtParam tool (https://web.expasy.org/protparam/) assuming all cysteines to be in the reduced form (ε = 1490 for AcpP; ε = 28880 for Sfp).

Sequence Analysis

Sequences for Camelina ACP proteins were originally obtained from Nguyen and colleagues (Nguyen et al., 2013). All listed sequences were downloaded from National Center for Biotechnology Information.

Protein Assay and SDS-PAGE

Protein Assay Reagent (Bio-Rad) was used for quantification of biological protein extractions with BSA as the protein standard. Mini-PROTEAN Tris-Tricine precast gels, 4 to 15% and 16.5% (w/v) were used for SDS-PAGE analysis. Proteins were visualized by Coomassie Brilliant Blue R 250 staining, imaged with Gel Doc EZ system and Image Lab 6.0.1 software (Bio-Rad); reaction efficiencies were analyzed by densitometry using ImageJ (Fiji version 2.0.0).

Acyl-ACP Preparation

Acyl-ACPs were enriched using TCA precipitation. In brief, fresh plant tissue from full, green seed 61 d after sowing (DAS) and young leaves (28 DAS) from the top of the plant was powdered in the presence of liquid nitrogen. Five percent TCA (w/w) was added to ∼50 mg of fresh tissue, and the solution was vortexed. The protein was pelleted with centrifugation at 21,000g for 10 min at 4°C. The supernatant was removed, and the process was repeated with precipitation in 1% (w/w) TCA. Next, the pellet was resuspended in 50 mM MOPS buffer, pH 7.5, incubated on ice for 1 h, and then centrifuged at 21,000g for 10 min at 4°C to remove cellular debris. Supernatant was collected and filtered, and TCA was added to a final concentration of 10% (w/w). The sample was frozen at –80°C for 1 h or overnight. After thawing on ice, the pellet was recovered by centrifugation at 21,000g for 10 min at 4°C and rinsed with 1% (w/w) TCA. A final centrifugation at 21,000g for 10 min at 4°C ensued prior to dissolving in a minimal volume of 50 mM MOPS buffer, pH 7.5.

Asp-N Protease Digestion

Acyl-ACP protein levels were quantified before samples were digested with endoproteinase Asp-N (Sigma-Aldrich) at 1:20 (enzyme:acyl-ACP, w/w), and incubated at 37°C overnight. Methanol was then added to a final concentration of 50% (v/v) to stop enzymatic digestion.

Assessing ACP Losses during Initial Development of the Protein Purification Steps

To gauge individual losses of ACPs during processing steps (although not crucial to the isotope dilution-based quantification), analyte acyl-ACPs from plant biomass were spiked with a 15:0-ACP standard before and during extraction steps as part of the initial development of the method. Recovery of a 15:0-ACP standard did not indicate a pattern in losses and was not specific to a particular step or changes in TCA concentration from 25 to 1% (w/w; Supplemental Tables 6 and 7).

Liquid Chromatography–Mass Spectrometry Conditions

The Asp-N proteolytic digestion products of acyl-ACPs were analyzed using a liquid chromatography tandem triple quadrupole mass spectrometer (QTRAP 6500 LC/MS/MS system, SCIEX). Analyst software (SCIEX version 1.6) was used to collect and analyze the data. The acyl-4-phophopantetheine-(DSL) tripeptides were first separated on a reversed phase column (Discovery BIO Wide Pore C18, Sigma-Aldrich; 10 cm, 2.1 mm, 3 μm) using solvent A (acetonitrile/10 mM ammonium formate and formic acid, pH 3.5, 10/90 [v/v]) and solvent B (acetonitrile/10 mM ammonium formate and formic acid, pH 3.5, 90/10 [v/v]). A flow rate of 0.2 mL/min was used with a gradient program: 0% B (100% A), at 4 min change to 10% B, then to 100% B at 12 min with a hold for an additional 5 min, then changed back to 0% B at 18 min and stopped at 27 min. The MS/MS spectrometry data were collected in positive ion mode. The curtain gas was set to 30 psi, the ion spray voltage was 4.5 kV, the source temperature was 400°C, the nebulizing gas was 30 psi, and the focusing gas was at 30 psi. The digested acyl-ACP molecules were identified using MRM mode. The masses for the precursor ions and product ions were calculated and an MRM list was generated. DP and CE were optimized by changing the values stepwise with standards or plant samples. The MRM masses, DP, and CE for routine analysis are recorded in Figure 3C, and the values for fatty acid elongation intermediates including 3-hydroxyacyl-ACPs and 2,3-trans-enoyl-ACPs are presented in Supplemental Table 2.

Optimization Conditions for Sfp Reactions

Optimization of Sfp reactions comprised of varying 50 mM MOPS from pH 6.5 to 7.5, DTT or TCEP from 0 to 5 mM, MgCl₂ from 5 to 20 mM, MnCl₂ from 5 to 20 mM, acyl- or BODIPY-FL-N-(2-aminoethyl)maleimide-S-CoA (BODIPY-CoA) from 150 to 500 µM, Sfp from 1 to 20 µM, and apo-ACP from 70 to 250 µM was performed at room, reduced temperature (4°C), or elevated temperature (37°C) for 1 h to 1 week.

BODIPY-CoA Chemical Synthesis

Chemical synthesis of the fluorophore BODIPY-CoA was accomplished by reacting 2.5 mg/mL BODIPY with 2.5 mg/mL CoA, tri-lithium in 100 mM MES, 100 mM Mg(OAC)₂, and 15% (v/v) DMSO at 4°C in the dark for 20 min. The reaction efficiency was assessed by thin layer chromatography. Unreacted dye was removed by extracting with ethyl acetate three times.

Preparation of Isotopically Labeled Acyl-ACP Standards

¹⁵N-Labeled acyl-ACP standards were synthesized enzymatically from ¹⁵N apo-ACP (overexpression in E. coli) with acyl-CoA substrates using Sfp from B. subtilis overexpressed in E. coli. Reactions performed in 50 mM MOPS, pH 6.5, 4 mM TCEP, 100 to 250 µM apo-ACP, 300 to 500 µM acyl-CoA, 10% DMSO, 1% Tween 20, 10 mM MgCl₂, 10 mM MnCl₂, and 25 µM Sfp were incubated for 3 h at 37°C.

Synthesized acyl-ACP standards were purified by TCA precipitation, and the reaction yields were determined. TCA was added to the reaction mixture to a final concentration of 5% (w/w) and the solution vortexed. The acyl-ACP was then pelleted by centrifugation at 21,000g at 4°C, and the supernatant discarded and washed with 1% (w/w) TCA followed by centrifugation. Pellets were resuspended in MOPS, pH 7.5, prior to use as an internal standard. The yield of each acyl-ACP standard was determined by SDS-PAGE on 16.5% (w/v) Tris-Tricine gels and confirmed by LC-MS/MS.

Isotope Dilution-Based Quantitation of ACPs in Biological Samples

Acyl-ACPs from developing seed and leaf tissue were quantified using internal standards and an isotope dilution-based approach. Biological replicates consisted of multiple aliquots from pooled seed or leaf tissue from several plants. Unlabeled standards were serially diluted with a constant concentration of ¹⁵N₃ acyl-ACP standards. The area ratios of unlabeled and labeled peaks were plotted against the concentration of the unlabeled standards in order to construct standard curves. Acyl-ACPs from unlabeled Camelina were spiked with ¹⁵N₃-labeled ACP standards and then prepared and analyzed as described above with quantities calculated using the aforementioned standard curves. All peak integration was performed using Analyst 1.6 software (SCIEX). Standard curves and quantitative calculations were performed using peak areas exported to Microsoft Excel. The limits of detection and quantification were obtained by first serial diluting standards until the peak area stopped decreasing. Next, the sd of the lowest concentration still resulting in a change in peak area was multiplied by 3 (for limits of detection [LODs]) and 10 (for limits of quantification [LOQs]), and divided by the slope of the calibration curve, to calculate LOD and LOQ.

Supplemental Data

• Supplemental Figure 1. Lowering MgCl₂ and MnCl₂ levels decreases 18:0-ACP synthesis by Sfp.

• Supplemental Figure 2. Solubilization of acyl-CoA reactants using DMSO and TWEEN-20 increases reaction efficiency.

• Supplemental Figure 3. Acyl-ACP Standards are quantitatively recovered by trichloroacetic acid (TCA) precipitation.

• Supplemental Figure 4. Complete digestion of acyl-ACP standards.

• Supplemental Figure 5. Calibration curves of acyl-ACPs.

• Supplemental Figure 6. Retention time for the 18:1-ACP elongation intermediate (i.e., C18-enoyl-ACP or 2,3-trans-octadecenoyl-ACP).

• Supplemental Figure 7. Plasmid design for ACP and Sfp transferase overexpression.

• Supplemental Table 1. Acyl-ACP synthesis reaction.

• Supplemental Table 2. Multiple-reaction monitoring (MRM) list for acyl-ACP elongation intermediates.

• Supplemental Table 3. Retention time of acyl-ACP elongation intermediates.

• Supplemental Table 4. Primers used for plasmid construction and sequencing.

• Supplemental Table 5. Overexpressed proteins used in this study.

• Supplemental Table 6. 15:0-ACP recovery.

• Supplemental Table 7. Recovery of 15:0-ACP with different TCA concentrations.

DIVE Curated Terms

The following phenotypic, genotypic, and functional terms are of significance to the work described in this paper: