Enzymatic synthesis and characterization of chlorophyllide derivatives as possible internal standards for pigment chromatographic analysis

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Enzymatic synthesis and characterization of chlorophyllide derivatives as possible internal standards for pigment chromatographic analysis Antonio Gavalás-Oleaa,*, Noelia Sanza, Pilar Rioboóa, José Luis Garridoa, Belén Vazb a

IIM-CSIC, Av. Eduardo Cabello 6, Vigo 36208, Spain Department of Organic Chemistry, Biomedical Research Center (CINBIO), and Southern Galicia Institute of Health Research (IISSG), Universidade de Vigo, 36310 Vigo, Spain

b

A R T I C LE I N FO

A B S T R A C T

Keywords: Chlorophyllide sters Internal Standard Pigment analysis Chromatography

In this article the chlorophyllase activity of Dunaliella salina has been employed to generate different chlorophyllide a and b esters that could potentially be used as internal standards in pigment analysis. Chlorophyllide a (8’-hydroxyoctyl) ester was selected due its adequate chromatographic and spectral properties and was fully characterized by UV-Vis, ESI-MS and NMR. An easy room temperature procedure for its synthesis is described. Attachment

1. Introduction The analysis of photosynthetic pigments, chlorophylls (Chls) and carotenoids, by liquid chromatography has become an essential tool in studies of physiology (e.g. in the characterization of the photosynthetic apparatus) and ecology (e.g. in the distribution of phytoplankton populations in natural waters) of photoautotrophic organisms [1–3]. The accuracy of the quantitative determination of pigments relies on the correct calibration of the chromatographic equipment by either internal (IS) or external (ES) standard methods [4,5]. Both calibration methods require a set of pure pigment standards. In the ES method, the chromatographic system is first calibrated with standards that contain the same pigments actually found in the sample. The peak heights or areas are correlated with the mass of the corresponding pigment injected to yield response factors for each pigment. These response factors are then used to calculate the mass of pigment, analyzed in separated injections, from the corresponding peak areas measured in the samples [5]. The IS method accounts for the effects of sample handling (dilution, evaporation, etc…) and for variations in the response of the chromatographic detectors [5], as they affect equally both the sample pigments and the IS. But these advantages are only really effective if the IS meets a series of strict requirements: it must be similar in analytical behavior (i.e. should be chemically similar) to the compounds of interest, but never present in the samples; it should elute completely resolved from other compounds in the sample; it must be stable and unreactive towards any other compound implied in the analysis (sample components,

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eluents, etc…) and, ideally, be readily available in pure form [5]. It has to be considered that the use of an IS constitutes also a source of uncertainty compared to the ES calibration, as it requires additional quantitative liquid handling and implies the measurement, for each compound to be determined, of two peak sizes instead of one [5]. Due to the different chemical nature of chlorophylls and carotenoids, an IS of general use for pigment analysis is difficult to find. Vitamin E acetate [6] has been proposed for such purpose as it is cheap, commercially available and, as it absorbs in the ultraviolet (UV) range (222 nm, far from the pigment absorption in the visible range of the spectrum), the co-elution with pigments is not an issue. However, although vitamin E acetate can be an excellent IS for correcting extraction volumes or injection errors, it would not be useful in the case of detection problems. In fact, this case can be especially important if the detector employs different lamps for UV and visible ranges of the spectrum, as any malfunction of the visible lamp used for pigment detection would not affect the response of vitamin E acetate, detected by the UV lamp. In addition, a single wavelength detector cannot be used if vitamin E acetate is selected as IS, while in the case of working with a double wavelength detector, this would lose its capacity of selectively detect some pigments (for example setting the wavelengths at 440 and 665 nm), as one of the detection channels should be reserved specifically for the IS. Furthermore, the use of vitamin E acetate is completely hampered in high performance liquid chromatography (HPLC) methods that employ UV-absorbing solvents, such as ethyl acetate or acetone. A different approach consists of using a compound absorbing in the

Corresponding author. E-mail address: [email protected] (A. Gavalás-Olea).

https://doi.org/10.1016/j.algal.2019.101688 Received 26 March 2019; Received in revised form 30 September 2019; Accepted 1 October 2019 2211-9264/ © 2019 Elsevier B.V. All rights reserved.

Please cite this article as: Antonio Gavalás-Olea, et al., Algal Research, https://doi.org/10.1016/j.algal.2019.101688

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checked daily (a must with any IS) by a short chromatographic analysis (the chromatographic run can be stopped soon after the IS appears). When the working solution is expended or degraded, a new solution can be taken from the freezer, checked and employed. If needed, these standards could be repurified by preparative HPLC. Chlorophyllide esters can be easily prepared by enzyme catalyzed transesterification using chlorophyllase [14]. Chlorophyllase (chlorophyll-chlorophyllide-hydrolase, EC 3.1.1.14) is an intrinsic membrane glycoprotein enzyme present in green plants and some unicellular marine algae [15–18] whose activity catalyzes two types of reaction: hydrolysis and transesterification [19]. Chlorophyllase can hydrolyze phytol esterified to the propionic acid side chain on C7 of the porphyrin ring of chlorophylls and pheophytins (and various derivatives of these compounds) to render the acidic chlorophyllides and pheophorbides [14]. In addition, it can catalyze transesterification reactions that can result in the incorporation of a variety of side chains to the porphyrin ring [20]. Importantly, this replacement of phytol by different alcohols is found to have no effect on the chromophore part of the molecule, thus, keeping the same value of the molar extinction coefficients in different solvents, and, thus, making them quantifiable compounds [21]. The enzyme chlorophyllase retains its activity in either aqueousorganic media prepared with water miscible solvents (acetone, low molecular weight alcohols) or in heterogenous biphasic systems [22] with organic solvents that facilitate the solubility of long chain alcohols. This capacity allows the design of adequate systems in which alcohols of different lengths could be employed for the synthesis of a collection of chlorophyllide esters showing distinctive chromatographic retention. As a result, specific chlorophyll derivatives could be tailor produced to elute in specific zones of the chromatogram where no other pigments do. In a survey for chlorophyllase activity in marine phytoplankton, the enzyme was detected in many eukaryotic algae [23] from both the red and the green evolutive lineages. With the purpose of synthesizing different chlorophyllides esters, green algae are especially useful as they possess both Chl a and Chl b and, thus, the same reaction can be employed to produce both Chlide a and Chlide b derivatives. Among chlorophytes, members of the genus Dunaliella showed high enzyme activities [23]. This paper shows the capacity of D. salina cell suspensions in either aqueous:organic or non-aqueous media to produce Chlide a and Chlide b esters (that can be used as IS in liquid chromatography of pigments) from its own native Chl a and Chl b in a simple and fast procedure. With no need of extraction or purification of the enzyme, the reaction is completed in about two hours. Chlorophyllide a (8′-hydroxyoctyl) ester obtained by this methodology features an adequate retention in two different chromatographic systems, and, therefore, is proposed as a possible IS for the HPLC analysis of chlorophylls.

range of visible wavelengths at which both chlorophylls and carotenoids absorb (420–460 nm) and employing it as a unique IS for both types of pigments (chlorohylls and carotenoids). However, as these compounds belong to different chemical groups, the use of two standards, one with properties similar to carotenoids, and a chlorophyll analogue would be preferred, so ideally both ISs could be added to the samples of the study. For carotenoids, the IS should be a natural or synthetic lipophilic compound strongly absorbing at 430–450 nm (preferably another carotenoid, either natural or synthetic). The use of carotenoids of terrestrial origin (β-apo-8′-carotenal, capsanthin) is common in the analysis of algal pigments [5–7]. The carotenoid canthaxanthin, although sometimes occurring in certain environments, shows limited distribution in oceanic waters and it has been successfully used with such samples [6]. The general lack of chlorophyll analogues with adequate chromatographic and spectral properties, plus satisfactory storage stability, purity, and commercial availability [8] led to the proposal of synthetic compounds as IS. Bohn and Walczyk [8] introduced the use of Znphthalocyanine, but its differences in absorption range with natural chlorophylls and its incompatibility with 90% aqueous acetone (a typical solvent for pigment extraction [9]) has hindered its use as a general standard. Similarly the use of the colorant Fast Green FCF, with marked spectral differences relative to natural chlorophylls, compromised the accuracy of their determination [10]. Several demetallated porphyrins and pheophorbides were tested as ISs. However, despite showing an adequate fluorescence, their weak absorbance at 410–460 nm, poor chromatographic behavior or lack of stability led to their rejection [11]. The synthesis of pigments with the desired properties (retention time, chemical stability and spectroscopic properties) from readily available natural chlorophylls as starting material has been proposed as an alternative. Mantoura and Repeta [11] informed about a method for the synthesis of a Zn derivative of pyropheophorbide a from Chl a using a set of reactions that included demetallation, hydrolysis of the phytol ester, pyrolysis of the 10-carboxymethyl ester, esterification of the propionic acid residue and transmetallation. Unfortunately, the detailed procedure for that synthesis was never published. Nevertheless, the complex set of specific reactions required for this synthesis makes their transfer to general laboratories impractical. Even though many new synthetic porphyrins and phtalocyanin dyes have become commercially available in recent years (Sigma-Aldrich, Porphychem), a systematic study of their spectral and chromatographic properties, as related with chlorophyll analysis, is still needed. Surprisingly, chlorophyllide (Chlide) esters sharing the tetrapyrrolic part of the molecule with chlorophylls and, thus, keeping identical spectral (absorption and fluorescence) properties, but differing in the lateral ester residue, have not been considered previously as possible ISs. This fact could be explained by their inherent moderate instability, as chlorophylls tend to epimerize or to allomerize in the presence of oxygen, as well as the acid lability of Mg2+ complexes, prone to demetallate under acidic conditions [12]. However, these drawbacks can also be attributed to any chlorophyll being analyzed; likewise the chlorophyll standards needed in either ES or IS calibration procedure. Nevertheless, with correct laboratory practice, and adopting the general precautions recommended for working with pigments (subdued light, cold solvents, avoidance of traces of acids or bases and storage under inert gas), these synthetic chlorophyllide esters different from phytol, could be considered an alternative as internal standards. The stability of these chlorophyll analogues should be similar to that of natural chlorophylls in the samples or commercial chlorophyll standards. Acetonic solutions of Chl a are stable when stored at -15 °C, with degradation rates in the order of -0.2% d−1 [13]. A quantitative IS solution in 90% acetone can be distributed in several pieces of a glassware proven to prevent evaporation and stored deep frozen. One of these solutions can be used as working solution, and its integrity can be

2. Materials and methods 2.1. Algal cultures and extraction procedure Dunaliella salina was obtained from the Culture Collection of Marine Microalgae, Instituto de Ciencias Marinas de Andalucía, Cádiz, Spain; cultures were grown at 15 °C in L1 enriched seawater medium [24], under 12:12 light/dark cycles with an irradiance of 100 μmol photons m−2s-1 during the light period. Cell counting was performed using a Beckman Coulter Counter (Nyon, Switzerland). Routinely, triplicated aliquots of 20 mL of culture (approximately 2*106 cells mL-1) were dosed in Teflon-lined screw capped tubes and centrifuged (Heraeus Biofugue-Stratos, Hanau, Germany) at 4500 rpm at 4 °C for 15 min. The supernatant was discarded and the pellets drained during 10 min in darkness by placing the tubes upside down on several sheets of filter paper to remove excess water. All cell harvesting and extraction operations were done under subdued light. 2

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equilibration time of 2 min in initial conditions was allowed between injections in both methods. Methanol and ethanol, both HPLC grade, were obtained from Panreac and Merck respectively. Ammonium acetate was analytical grade from Carlo Erba (Sabadell, Spain). Samples were mixed with Milli-Q water to avoid peak distortion [27,28] by either manually adding 0.4 mL of Milli-Q to 1 mL of each sample extract immediately before injection (200 μL), or by employing an automatic injection sequence in which 74 μL water, 130 μL sample and 37 μL water were consecutively placed in the injection loop. Pigments were identified by co-chromatography with authentic standards obtained from SCOR reference cultures [29] and diode-array spectroscopy (see Zapata et al. [25]). After checking for peak purity, spectral information was compared with a library of Chl and carotenoid spectra from pigments extracts prepared from phytoplankton cultures [30,31].

To determine pigment content of the culture in each experimental condition, cell pellets were extracted in 10 mL of 90% aqueous acetone. The tubes were placed in an ultrasonic bath with ice and water and sonicated for 5 min. Tubes were subsequently centrifuged for 5 min at 3500 x g at 4 °C to remove cell debris. Extracts were filtered through 13 mm diameter syringe filters (MS PTFE, 0.22 μm pore size, hydrophilic PTFE, Membrane Solutions, Plano, Texas, USA) previous to be injected in the HPLC system. 2.2. Transesterification The retention time of the derivatives were studied using different set of alcohols: linear saturated aliphatic alcohols (methanol, ethanol, propan-1-ol, butan-1-ol, pentan-1-ol, hexan-1-ol, octan-1-ol, decan-1-ol and dodecan-1-ol), branched chain saturated aliphatic alcohols (propan-2-ol, butan-2-ol) and linear saturated aliphatic primary diols (butane-1,4-diol, hexane-1,6-diol, octane-1,8-diol, decane-1,10-diol). Control reaction media were prepared with 1:1 mixtures of acetone and water. Incubation media for transesterification (2 mL) were prepared depending on the alcohol solubility: a) 1:1 mixtures of water and one of the following short chain alcohols (up to 4 carbon atoms: methanol, ethanol, propan-1-ol, propan-2-ol, butan-1-ol or butan-2-ol); b) 1:1 mixtures of hexane and one of the following long chain alcohols (pentan-1-ol, hexan-1-ol, octan-1-ol, decan-1-ol and dodecan-1-ol), and c) 1:1 mixtures of water and one of the following acetonic solutions of diols (50 mg mL−1 for butane-1,4-diol, hexane-1,6-diol and octane-1,8diol and 0.3 mg mL−1 for decane-1,10-diol). Samples (triplicated cell pellets resulting from the centrifugation of 20 mL of the culture as described in section 2.1) were gently resuspended in 2 mL each of the reaction media (as described above) and the mixtures were incubated two hours at room temperature (25 °C) in the dark, with brief periods of gentle agitation each 15 min. The reaction was stopped, and the pigments extracted in the same tube, by adding 8 mL acetone to incubation mixtures a) and c) or 7 mL acetone and 1 mL water to incubation mixtures b), to achieve 10 mL of 10% aqueous pigment extracts. Reagent-grade chemicals were employed: acetone, methanol, hexane, propan-2-ol, butan-1-ol, octan-1-ol from Panreac (Castellar del Vallés, Spain); ethanol and decan-1-ol from Merck (Darmstadt, Germany); hexan-1-ol, dodecan-1-ol, propan-1-ol, pentan-1-ol, butan-2ol, butane-1,4-diol, hexane-1,6-diol; octane-1,8-diol and decane-1,10diol from Sigma-Aldrich (St. Louis, Missouri, USA) (please consider that some providers still use the old nomenclature for alcohols, e.g., 1,8octanediol).

2.4. Internal standard purification and stability A large volume of extracts (in 90% aqueous acetone) containing the derivative pigment was prepurified using a solid phase separation cartridge (Sep-Pak C18 Vac 35cc, 10 g, Waters). After conditioning the cartridge with several volumes of absolute acetone followed by 40% aqueous acetone, the extract diluted to 45% aqueous acetone was eluted through the cartridge allowing the retention of whole pigment extract. Pigments were eluted with aqueous acetone solutions with increasing proportions (+5%) of acetone. A fraction of Chlide a (8′-hydroxyoctyl) ester was obtained by eluting with 70% aqueous acetone. The fraction was concentrated using a similar procedure, in which pigment was retained in 45% acetone with a solid phase separation cartridge (Sep-Pak C18 Plus, Waters) and eluted in 100% acetone. A preparative HPLC equipment consisting of a Waters 1525 Binary HPLC Pump and a Waters 2489 UV/visible detector (set at 440 and 665 nm) with a Luna C8 column (250 x 10 mm, 5 μm) from Phenomenex (Madrid, Spain) was used for preparative chromatography. An isocratic method (90:10 methanol/water) at a flow rate of 5 mL min−1 during 15 min was employed. Pure compound of interest was isolated by collecting the chromatographic peak from several runs and was concentrated using a solid phase separation cartridge (as previously described), the cartridge was dried with N2 and the pigment was finally eluted with absolute acetone. For the study of stability of the selected IS with time, a solution of Chlide a (8′-hydroxyoctyl) ester (0.001 μmol/mL) in 90% acetone was aliquoted in 1 mL vials and stored under N2 at −20 °C and their purity checked by HPLC [25] every week during 50 days. 2.5. UV–vis spectrophotometry

2.3. HPLC pigment analysis UV-visible spectra (350–700 nm) of purified pigments were recorded in acetone using a V-650 Spectrophotometer (Jasco, Tokyo, Japan). To avoid evaporation of solvent, special optical glass cuvettes with Teflon caps were used. Spectrophotometer settings were as follows: bandwidth 2.0 nm, data interval 0.5 nm and scan speed 400 nm min−1.

Pigments were separated using a Waters Alliance HPLC System (Waters Corporation, Milford, Massachusetts, USA) consisting of a 2695 separations module, a Waters 996 photo-diode-array detector (PDA, 1.2 nm optical resolution). Pigment separation was performed using the HPLC method of Zapata et al. [25], with a reformulated mobile phase A [26]. The column was a C8 Waters Symmetry (150 × 4.6 mm, 3.5 μm particle size, 100 Å pore size). Flow rate was set at 1.0 mL min−1, and column temperature was fixed at 25 °C. Mobile phases were A) Methanol: Acetonitrile: 25 mM aqueous Pyridinium Acetate at pH = 5 (50:25:25; v:v:v) and B) Methanol: Acenotonitrile: Acetone (20:60:20, v:v:v), see gradient in Table S-1. Solvents were HPLC grade (Panreac) and pyridine was reagent grade (Merck). A volume of the proposed Chlide a (8′-hydroxyoctyl) ester solution was added to filters of natural samples from Atlantic Ocean just prior to the extraction. These extracts were then chromatographed with Sanz et al. [7] method using a ACE C18 PFP column (150 mm - 4.6 mm, 3 μm particle size; Advanced Chromatography Technologies, Aberdeen, UK) at 33 °C and a flow rate of 1 mL min−1. Mobile phases were A) Methanol: 225 mM Ammonium Acetate (82:18; v:v) and B) Ethanol, see gradient in Table S-2. A re-

2.6. Mass spectrometry High resolution positive ion mass spectra of Chlide a (8′-hydroxyoctyl) ester were obtained with a Thermo Scientific Dionex Ultimate 3000 High-Speed LC (Waltham, Massachusetts,USA) (column: Waters Acquity HSS C18, 1.8 μm, 150 x 2.1 mm; isocratic elution with 0.1 % formic acid in 9:1 methanol/water, flow rate 200 μL min−1) coupled to a Thermo Scientific Exactive mass spectrometer (Waltham, Massachusetts,USA), equipped with an Orbitrap mass analyzer and a HESI-II probe for electrospray ionization. Full scan and AIF (all ion fragmentation) scans were acquired. Mass calibration was performed with a mixture consisting of caffeine, MRFA tetrapeptide, and Ultramark 1621 (Thermo Scientific). All analyses were performed using 3

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the “balanced” automatic gain control (AGC) setting with a 200 ms maximum inject time. Data acquisition was carried out using Thermo Scientific Xcalibur 2.1. Optimal ion source and interface conditions consisted of a spray voltage of 4 kV, sheath gas flow of 25, auxiliary gas flow of 7, capillary temperature of 320 °C, In source CID 0.0 eV. For Full MS spectra conditions were as follows: Microscans 1; Resolution 70,000; AGC target 3e6; Maximum IT 200 ms; Scan range 120–1200 m/ z. In AIF mode: Microscans 1; Resolution 35,000; AGC target 3e6; Maximum IT 100 ms; NCE (normalized collision energy) 35.0; Scan range 120–1000 m/z.

reverse elution order in the case of the analogous derivatives of Chlide b (Table 1). As expected, the corresponding diols were less retained than its analogous primary alcohols (Fig. 2), due to the presence of a hydroxyl group at the end of the alkyl chain that increases their polarity and promotes a faster elution in the chromatogram (Fig. 2 and Table 1). The possibility of the formation of double esters (when diols were used) was discarded by HPLC with a modification of the method of Zapata et al. [25]: after the gradient was completed (min 38) the eluent was gradually changed to 100% acetone where it remained for further 60 min. No additional peak was observed in the prolonged method. Probably, the formation of these diesters was prevented by steric hindrance between the two chlorin macrocycles. Importantly, the derivatives of octane-1,8-diol and dodecan-1-ol elute at retention windows in which no other pigment is expected (Fig. 2 and Table 1). For those methods focused in the determination of Chls a and b (even losing the resolution of polar chlorophylls, [32]) an internal standard expected to elute in their vicinity can be more adequate. With that purpose, the use of dodecan-1-ol to synthetize the derivative in totally organic medium is proposed. For those more general methods intended for the determination of both polar and non polar chlorophylls, Chlide a (8′-hydroxyoctyl) ester results to be a more general use IS: it elutes more centrally in the chromatogram, in a wider zone free of other pigments and avoids the use of a more toxic organic solvent (hexane). The capacity of octane-1,8-diol derivatives as possible ISs was also tested in alternative general HPLC methods. Thus, the analysis of a natural sample using the method of Sanz et al. [7] provided a well resolved chromatogram where Chl a ester derived from octane-1,8-diol eluted separately from the natural pigments under study (Fig. 3).

2.7. Nuclear magnetic resonance 1D and 2D-NMR spectra were measured in deuterated tetrahydrofuran (THF-d8) by a Bruker Avance 600 MHz NMR spectrometer (Billerica, Massachusetts, USA) at room temperature with residual protic solvent as the internal reference. Chemical shifts (λ) are given in parts per million (ppm), and coupling constants (J) are given in Hertz (Hz). Assignments of 1H signals were obtained by the analysis of 1H-1H correlated spectroscopy (COSY) and nuclear overhauser effect spectroscopy (NOESY) correlations. 3. Results and discussion 3.1. HPLC pigment study The chromatographic analysis of the extracts obtained after the incubation of cell suspensions of D. salina in media containing different alcohols allowed the detection of peaks with spectra similar to that of Chl a and Chl b, which were tentatively identified as the corresponding Chlide a (Fig. 1) and b esters. Fig. 2 shows the distinctive positions of time elution for the different Chlide a and b esters. Additionally, noncoincident coelutions were observed for derivatives of both chlorophylls (Fig. 2 and Table 1). As expected, the chromatographic retention of the esters of aliphatic alcohols is governed by its chain length, with minimum retention of the esters corresponding to methyl Chlides a and b and maximum to the dodecyl esters (Table 1). The branched chain aliphatic alcohols tested, propan-2-ol and butan-2-ol, produced Chlide a esters slightly less retained relative to those of the corresponding primary alcohols, with a

3.2. Structural characterization of Chlide a (8′-hydroxyoctyl) ester As expected, the spectral absorption properties in the visible region (measured on line in the chromatogram with the PDA detector) were compatible with that of Chl a, suggesting that they share the same chromophore (Fig. 4a). The on-line spectra during the chromatographic run show differences that can be explained by the different nature of the solvent at the times of elution of both compounds (as it happens with Chlide a and Chl a as described in [31]). When purified samples of both pigments were dissolved in absolute acetone and their respective

Fig. 1. Different alcohol derivatives of Chlide a. 4

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Fig. 2. Chromatograms of the DHI mixed pigments standards with Zapata et al. [25] method showing the elution position of different straight chain alcohol derivatives of Chlide a (a) and Chlide b (b). Red dashed lines show the retention position of chlorophyllide esters from diols (peak numbers as in Table 1) and blue dashed lines show esters derived from primary alcohols. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

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Table 1 Retention time of pigment and ester derivatives in Zapata et al. method [25]. Orange cells represent coelution with other pigments while green cells mean no coelution. Abbreviations in table are as follows: Chl c3 (chlorophyll c3), MgDVP (Magnesium 2,4-divinylpheoporphyrin a5 monomethyl ester), Chl c2 (chlorophyll c2), Chl b (chlorophyll b), DVChl b (divinyl chlorophyll b), DVChl a (divinyl chlorophyll a), Chl c2- MGDG (chlorophyll c2-monogalactosyldiacylglyceride ester [18:4/ 14:0]), Chl a (chlorophyll a).

Fig. 3. Chromatogram of a natural sample from Atlantic Ocean (N 33° 1′ 16.32′' O 12° 14′ 26.016′') obtained by Sanz et al. method [7]. Peak identification = 1: Chl c3; 2: Peridinin; 3: Chl c2; 4: MgDVP; 5: 19′-Butanoyloxyfucoxanthin; 6: Fucoxanthin; 7: 9′-cis-Neoxanthin; 8: 19′-Hexanoyloxyfucoxanthin; 9: Prasinoxanthin; 10: Violaxanthin; 11: Chlide a (8′hydroxyoctyl) ester; 12: Diadinoxanthin; 13: Astaxanthin; 14: Anteraxanthin; 15: Alloxanthin; 16: Zeaxanthin; 17: β-apo-8′carotenal; 18: DVChl b; 19: Chl b; 20: DVChl b epimer; 21: Chl b epimer; 22: DVChl a; 23: Chl a; 24: Unknown carotenoid of Osterococcus; 25: DVChl a epimer; 26: Chl a epimer; 27: β,εcarotene; 28: Pheophytin a; x: carotenoid-like spectrum; and y: chlorophyl-like spectrum.

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Fig. 4. a) Visible spectra and Absorption maxima in mobile phase of Chl a (blue solid line, retention time at 33.51 min) and Chlide a (8′- hydroxyoctyl) ester (red dashed line, retention time at 15.59 min, Zapata et al. method [25]). Spectra were normalized at 665 nm. b) Visible spectra and absorption maxima of Chl a (blue solid line) and Chlide a (8′- hydroxyoctyl) ester (red dashed line) in acetone. Spectra were normalized at 661.5 nm. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

Fig. 5. Full ESI-MS spectrum of octane-1,8-diol derivative and its structure.

spectra recorded in the spectrophotometer, an almost complete match of the spectra was obtained, with exact coincidence of their absorption maxima (Fig. 4b). High resolution ESI analysis (Fig. 5) of the octane-1,8-diol derivative provided an abundant [M+H]+ peak (at m/z 743.3644) in accordance with the neutral formula C43H51MgN4O6, which correlates

well with the structure proposed (m/z 743.366 calculated monoisotopic mass, deviation 2.02 ppm). This neutral formula features eight carbons (together with the corresponding hydrogens) and an additional oxygen atom in comparison to the corresponding structure of Chlide a (C35H34MgN4O5). Likewise, sodiated (m/z 765.3448) and potasiated ions (m/z 781.3190) were in accordance with calculated monoisotopic 7

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Fig. 6. Low field 1H-NMR spectrum of in Chlide a (8′-hydroxyoctyl) ester in THF-d8 at room temperature identifying corresponding signals of chlorin macrocycle. Table 2 1 H-NMR signal assignments and NOE correlations for Chlide a (8′-hydroxyoctyl) ester in THF-d8. Chl a assignments in acetone- d6 taken from Hynninen et al. [37]. Signal numbers are according to Fig. 7. Assignment

Chemical shift δ (ppm)

Integration and Multiplicity (JH-H (Hz))

Protons having NOE correlations

Chl a chemical shift δ (ppm)

Chl a (phytol not included) integration and Multiplicity (JH-H (Hz))

H10 H5 H20 H31 H32-trans H132 H32-cis H18 H17 H1’ H81 MeO132 Me121 H8’ Me21 Me71 OH8’ H172a H172b H171a H171b Me181 Me82 H2’ H3’, H7’ H4’, H5’, H6’

9.62 9.37 8.45 8.10 6.21 6.20 5.94 4.52 4.16 3.97 3.82 3.73 3.61 3.37 3.35 3.30 3.20 2.63 2.62 2.43 2.14 1.77 1.73 1.49 1.39 1.25

1H, 1H, 1H, 2H, 1H, 1H, 1H, 1H, 1H, 1H, 2H, 3H, 3H, 2H, 1H, 3H, 1H, 1H, 1H, 1H, 1H, 3H, 3H, 2H, 4H, 6H,

H81, Me82, Me121 Me71, H31 Me21, Me181, H18 H32−cis, H5 H32−cis, Me2, H5 H171, H17 H31, H32−trans Me181, H17, H20 H171, H18, H132, Me181 H2’ Me71, H10, Me82 – H10 – H32−trans, H20 H5, H81 – H132, H17 H132, H17 – – H20, H17, H18 Me71, H81 H1’ – –

9.57 9.27 8.40 8.04 6.14 6.06-6.11 5.92 4.18 4.06 – 3.67 3.72 3.49 – 3.24 3.18 – 2.37-1.96 2.37-1.96 2.37-1.96 2.37-1.96 1.66 1.61 – – –

1H, 1H, 1H, 2H, 1H, 1H, 1H, 1H, 1H, – 2H, 3H, 3H, – 1H, 3H, – 1H, 1H, 1H, 1H, 3H, 3H, – – –

‒ 4.13 ‒ 3.94 ‒ 3.79

‒ 3.33

‒ ‒ ‒ ‒ ‒ ‒

2.59 2.57 2.38 2.07 1.73 1.70

‒ 1.33 ‒ 1.18

s s s dd (17.8 and 11.5) d (17.8) s d (11.5) c (7.4) m m c (7.5) s s m s s t (5.2) m m m m m m app t (7.1) app quint (6.9) m

s s s dd (18 and 12) dd (18 and 1) s dd (12 and 1) q (7 and 7) q (7 and 7) q (7) s s s s m m m m d (7) t (7)

NMR spectroscopy using THF-d8 as deuterated solvent. This solution of the putative Chlide a (8′- hydroxyoctyl) ester was degassed with freeze–thaw cycles under Ar atmosphere and subdued light in order to avoid pigment degradation. Analysis of 1H-NMR spectrum showed meso-CH signals matching those previously described [34–36] in the lowest field region (between δ: 9.7-8.4). These signals were assigned to C10, C5 and C20 in order of appearance from low to high field (Fig. 6, Table 2). The following spin system featuring three protons corresponds to a peripheral vinyl group at position C3 [C31: δ =8.10 ppm (dd, J = 17.8 and 11.5 Hz, 1 H) and the terminal vinyl protons at C32: δ = 6.21 (d, J =17.8 Hz, 1H-trans) and δ = 5.94 (d, J =11.5 Hz, 1H-cis)]. Overlapping this spin system a characteristic singlet was observed, coming from the proton at the stereogenic centre at C132 (δ =6.20 ppm). This signal showed NOESY correlations with protons at C17 and C171 confirming the spatial proximity between both moieties (Fig. 7, Table 2). All these signals at low-field are in concordance with previously published resonance data

masses as well (m/z 765.3448, deviation 3.98 ppm; and m/z 781.3190 deviation 3.56 ppm, respectively). Further tandem MS studies (Figure S-1) revealed two peaks consistent with the fragmentation of the methoxycarbonyl substituent at C132, probably promoted by the benzylic character of the positive charges generated in the MS/MS experiment. Thus, a loss of 32 Da from the formal loss of H3COH is observed, resulting in an ion mass of m/z 711.3377, together with a 60 Da mass difference, from the loss of HCOOCH3, providing an ion peak of m/z 683.3437. In contrast, no fragment from the loss of the octane-1,8-diol was observed, as no aromatic effect is observed on the fragmentation pattern at C17 position. This lack of fragment ions resulting from the loss of saturated esterifying alcohols (in contrast to the common loss of phytol or farnesol) has also been observed in the mass spectra of secondary homologues of bacteriochlorophylls, esterified by straight and branched chain saturated alcohols [33]. Additional structural information was obtained from 1D- and 2D8

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directly affected by the aromatic ring current of the porphyrin, appeared close to the methoxy group at C132 as 3H-integrating singlets: MeO132 at δ = 3.73 followed by Me121 at δ = 3.61, Me21 at δ = 3.35 and Me71 at δ = 3.30. At higher field, signals of the methyl groups distant from the aromatic ring were identified only by COSY, as they overlap with the residual protic solvent signal at 1.73 ppm. Thus, multiplets at δ = 1.77‒1.73 (3 H) and δ = 1.73‒1.70 (3 H) were assigned to Me181 and Me82 respectively. With the aid of COSY spectrum, four additional signals belonging to the methylenes of the propionic acid appeared in the high field region (H171 and H172 between δ = 2.63-2.07), showing differentiated chemical shifts for each of the diastereotopic protons of this moiety. Finally, the most shielded chemical shifts corresponding to the central region of the esterified 8′hydroxyalcohol where assigned to the coincident H3′, 4′, 5′, 6′ identified by COSY correlations. 2D-NMR (COSY and NOESY, Figures S-4, S-5, S-6) spectroscopy confirmed the resonance assignments of H1-NMR spectrum. Importantly, NOE correlations revealed spatial proximity of methyl substituents around the porphyryn skeleton as shown in Fig. 7 and Table 2, which are in agreement with the characteristic substitution of Chl a. This data match previously correlation data published for Chl a [35–37].

Fig. 7. COSY and NOE correlations identified in Chlide a (8′-hydroxyoctyl) ester.

for Chl a [34–37]. The low intensity multiplets centered at 4.52 and 4.15 ppm could be assigned to the protons at positions C18 and C17 respectively, which in turn showed COSY correlations to the methyl group at C18 (∼1.74 ppm) and the methylene protons of the propionic acid side chain at C171 (∼2.43 ppm). In the same region of the spectra the first signals corresponding to the esterified alcohol could be identified, where H1’ appeared as a multiplet at δ = 3.97-3.94 ppm followed by the also deshielded signals proximal to the alcohol group at C8′, corresponding to H8′ (at δ = 3.37-3.33, 2H, m) and the hydroxyl group (8′OH, at δ = 3.20). Methyl substituents of porphyrin skeleton,

3.3. Incubation protocol and stability study Once the IS candidate (Chlide a (8′-hydroxyoctyl) ester) was established and characterized, different additional experiments were performed to optimize the variables that can affect the hydrolysis and esterification process. Several parameters influencing the transesterification with octane-1,8-diol were tested to set an optimized incubation protocol. Transesterification of Chlide a (8′-hydroxyoctyl) ester was

Fig. 8. Incubation time in octane-1,8-diol derivative experiments (triplicated) and chromatograms of Dunaliella salina [25] before (a) and after 2 h incubation with octane-1,8-diol (b). Abbreviations in figure are as follows: Chlide a (8′-hydroxyoctyl) ester (Chlorophyllide a (8′-hydroxyoctyl) ester), Chlide b (8′-hydroxyoctyl) ester (Chlorophyllide b (8′-hydroxyoctyl) ester), c-Neo (9′-cis- Neoxanthin), Viola (Violaxanthin), Anth (Antheraxanthin), Zea (Zeaxanthin), Lut (Lutein), Chl b (Chlorophyll b), Chl a (Chlorophyll a) βψ-Car (βψ-carotene), ββ-Car (ββ-Carotene). 9

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monitored after 0.5, 1, 2, 3, 4, 8, 12 and 24 h of incubation. The reaction resulted in an increased derivative production with time, reaching a plateau between 2 and 4 h after incubation (Fig. 8). The chlorophyllase assay protocol was set to 2 h, optimizing the time/derivative ratio (Fig. 8). 25 °C (room temperature) was adopted as the working temperature following Jeffrey & Hallegraeff [23], as it eliminates the need of a controlled temperature incubation system, thus, simplifying the protocol. Octane-1,8-diol concentration was a determining factor for establishing the optimal incubation conditions. Chlorophyll derivatives levels increased with the concentration of reactant in the reaction medium, reaching a maximum performance when saturation point was set to 0.1 g mL−1 (Data not shown). The purified compound was stored at −20 °C in 90% acetone to check its stability over time. The derivative and the formation of possible alteration products remained roughly at same levels, starting from the beginning of the sampling to a storage time of 50 days (Figure S-7). The degradation rate of Chlide a (8′-hydroxyoctyl) ester was 0.095 % d−1, a value lower than the degradation of Chl a in acetonic extracts described by Hooker et al. [13]. Nevertheless, the determination of the accurate concentration of Chlide a (8′-hydroxyoctyl) ester solutions previous to its addition as IS to the samples is recommended.

[3] S.W. Jeffrey, S.W. Wright, M. Zapata, Microalgal classes and their signature pigments, in: S. Roy, in: C.A. Llewellyn, E.S. Egeland, G. Johnsen (Eds.), Phytoplankton Pigments: Characterization, Chemotaxonomy and Applications in Oceanography, Cambridge University Press, Cambridge, U.K, 2011, pp. 1–77. [4] J. Asshauer, H. Ullner, Quantitative analysis in HPLC, in: H. Engelhardt (Ed.), Practice of High Performance Liquid Chromatography, Springer, Berlin Heidelberg, 1986, pp. 66–108. [5] L.R. Snyder, J.J. Kirkland, Quantitative and trace analyses, Introduction to Modern Liquid Chromatography, second edition, John Wiley and sons, New York, 1979, pp. 541–579. [6] R.R. Bidigare, L. Van Heukelem, C. Trees, Analysis of algal pigments by high-performance liquid chromatography, in: R.A. Andersen (Ed.), Algal Culturing Techniques, Phycological society of America, Burlington, Massachusetts, 2005, pp. 327–345. [7] N. Sanz, A. García-Blanco, A. Gavalás-Olea, P. Loures, J.L. Garrido, Phytoplankton pigment biomarkers: HPLC separation using a pentafluorophenyloctadecyl silica column, Methods Ecol. Evol. 6 (2015) 1199–1209. [8] T. Bohn, T. Walczyk, Determination of chlorophyll in plant samples by liquid chromatography using zinc-phthalocyanine as an internal standard, J. Chromatogr. A 1024 (2004) 123–128. [9] J.L. Pinckney, D.F. Millie, L. Van Heukelem, Update on filtration, storage and extraction solvents, in: S. Roy, C.A. Llewellyn, E.S. Egeland, G. Johnsen (Eds.), Phytoplankton Pigments: Characterization, Chemotaxonomy and Applications in Oceanography, Cambridge University Press, Cambridge, U.K, 2011, pp. 627–635. [10] S.C. Huang, C.F. Hung, W.B. Wu, B.H. Chen, Determination of chlorophylls and their derivatives in Gynostemma pentaphyllum Makino by liquid chromatographymass spectrometry, J. Pharm. Biomed. Anal. 48 (2008) 105–112. [11] R.F.C. Mantoura, D. Repeta, Calibration methods for HPLC, in: S.W. Jeffrey, R.F.C. Mantoura, S.W. Wright (Eds.), Phytoplakton Pigments in Oceanography, Unesco Publishing, Paris, France, 1997, pp. 407–428. [12] P.H. Hynninen, Chemistry of chlorophylls: modifications, in: H. Scheer (Ed.), Chlorophylls, CRC press. Taylor and Francis group, Boca Raton, Florida, 1991, pp. 145–209. [13] S.B. Hooker, L. Van Heukelem, C.S. Thomas, H. Claustre, J. Ras, R. Barlow, H. Sessions, L. Schlueter, J. Perl, C. Trees, V. Stuart, E. Head, L. Clementson, J. Fishwick, C. Llewellyn, J. Aiken, The Second SeaWiFS HPLC Analysis RoundRobin Experiment (SeaHARRE-2), NASA Tech. Memo, Goddard Sp. Flight Cent. 2005–21278, (2005) 112 pp. [14] R.F. McFeeters, Chlorophyllase, in: J.R. Whitaker, A.G.J. Voragen, D.W.S. Wong (Eds.), Handbook of Food Enzymology, Marcel Dekker, New York, USA, 2002, pp. 681–686. [15] J. Barrett, S.W. Jeffrey, Chlorophyllase and formation of an atypical chlorophyllide in marine algae, Plant Physiol. 39 (1964) 44–47. [16] J. Barrett, S.W. Jeffrey, A note on the occurrence of chlorophyllase in marine algae, J. Exp. Mar. Bio. Ecol. 7 (1971) 255–262. [17] W. Terpstra, Idenfication of Chlorophyllase as a Glycoprotein 126 North-Holl. Biomed. Press, 1981, pp. 231–235. [18] R. Suzuki, Y. Fujita, Chlorophyll decomposition in Skeletonema costatum: a problem in chlorophyll determination of water samples, Mar. Ecol. Prog. Ser. 28 (1986) 81–85. [19] S.I. Beale, Biosynthesis of photosynthetic pigments, in: N.R. Baker, J. Barber (Eds.), Chloroplast Biogenesis, Elsevier, Amsterdam, 1984, pp. 133–205. [20] T.J. Michalski, J.E. Hunt, C. Bradshaw, A.M. Wagner, J.R. Norris, J.J. Katz, EnzymeCatalyzed organic syntheses: Transesterification reactions of chlorophyll a, bacteriochlorophyll a, and derivatives with chlorophyllase, J. Am. Chem. Soc. 110 (1988) 5888–5891. [21] A.S. Holt, E.E. Jacobs, Spectroscopy of plant pigments. I. Ethyl Chlorophyllides a and b and their pheophorbides, Am. J. Bot. 41 (1954) 710–717. [22] A. Khamessan, S. Kermasha, P. Marsot, Effects of polar organic solvents on the biocatalysis of chlorophyllase in a biphasic organic system, Biosci. Biotechnol. Biochem. 58 (1994) 1947–1952. [23] S.W. Jeffrey, G.M. Hallegraeff, Chlorophyllase distribution in ten classes of phytoplankton: a problem for chlorophyll analysis, Mar. Ecol. Prog. Ser. 35 (1987) 293–304. [24] R.R.L. Guillard, P.E. Hargraves, Stichochrysis immobilis is a diatom, not a chrysophyte, Phycologia 32 (1993) 234–236. [25] M. Zapata, F. Rodríguez, J.L. Garrido, Separation of chlorophylls and carotenoids from marine phytoplankton: a new HPLC method using a reversed phase C8 column and pyridine-containing mobile phases, Mar. Ecol. Prog. Ser. 195 (2000) 29–45. [26] J.L. Garrido, S. Roy, The use of HPLC for the characterization of phytoplankton pigments, in: D.B. Stengel, S. Connan (Eds.), Natural Products from Marine Algae. Methods and Protocols, Springer, Humana Press, New York, 2015, pp. 241–275. [27] M. Zapata, J.L. Garrido, Influence of injection conditions in reversed-phase highperformance liquid of chromatography of chlorophylls and carotenoids, Chromatographia 31 (1991) 589–594. [28] M. Latasa, K. van Lenning, J.L. Garrido, R. Scharek, M. Estrada, F. Rodríguez, M. Zapata, Losses of chlorophylls and carotenoids in aqueous acetone and methanol extracts prepared for RPHPLC analysis of pigments, Chromatographia 53 (2001) 385–391. [29] S.W. Jeffrey, S.W. Wright, Qualitative and quantitative HPLC analysis of SCOR reference algal cultures, in: S.W. Jeffrey, R.F.C. Mantoura, S.W. Wright (Eds.), Phytoplakton Pigments in Oceanography, Unesco Publishing, Paris, France, 1997, pp. 343–360. [30] E.S. Egeland, Appendix C. Minimum identification criteria for phytoplankton pigments, in: S. Roy, C.A. Llewellyn, E.S. Egeland, G. Johnsen (Eds.), Phytoplankton Pigments: Characterization, Chemotaxonomy and Applications in Oceanography,

4. Conclusions The chlorophyllase activity of the microalga Dunaliella salina allowed a fast, easy and environmental friendly method to synthesize both chlorophyllide a and b esters from a wide variety of alcohols. One of them, Chlide a (8′-hydroxyoctyl) ester, has been proposed as a possible IS for HPLC pigment analysis. Briefly, a pellet of Dunaliella salina biomass was incubated with 50% aqueous-acetonic solution (containing octane-1,8-diol) during 2 h, after which time, the reaction was stopped and the pigments extracted with absolute acetone to a final 90% acetone proportion. The proposed IS, Chlide a (8′-hydroxyoctyl) ester, shows polarity characteristics that allows its elution in segments of the chromatogram free from other pigments in two different HPLC methods. In addition, it can be stored for up to 50 days, with good stability. Declaration of Competing Interest The authors declare that there is no conflict of interest. No conflicts, informed consent, human or animal rights are applicable for this work. Acknowledgments We thank Patricia Loures for her technical support. This research was co-funded by the Spanish Ministerio de Economía y Competitividad (MINECO) and FEDER through project CTM2012-32181. This work is appended as part of A. Gavalás-Olea DO*MAR Phd thesis (University of Vigo), who also acknowledge his doctoral FPI grant (BES-2013065752). 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.algal.2019.101688. References [1] C. Brunet, G. Johnsen, J. Lavaud, S. Roy, Pigments and photoacclimation processes, in: S. Roy, C.A. Llewellyn, E.S. Egeland, G. Johnsen (Eds.), Phytoplankton Pigments: Characterization, Chemotaxonomy and Applications in Oceanography, Cambridge University Press, Cambridge, U.K, 2011, pp. 445–471. [2] S.W. Jeffrey, Application of pigment methods to oceanography, in: S.W. Jeffrey, R.F.C. Mantoura, S.W. Wright (Eds.), Phytoplakton Pigments in Oceanography, Unesco Publishing, Paris, France, 1997, pp. 127–166.

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Cambridge University Press, Cambridge, U.K, 2011, pp. 650–652. [31] E.S. Egeland, Data sheets aiding identification of phytoplankton carotenoids and chlorophylls, in: S. Roy, C.A. Llewellyn, E.S. Egeland, G. Johnsen (Eds.), Phytoplankton Pigments: Characterization, Chemotaxonomy and Applications in Oceanography, Cambridge University Press, Cambridge, U.K, 2011, pp. 665–822. [32] S.W. Wright, S.W. Jeffrey, R.F.C. Mantoura, C.A. Llewellyn, T. Bjørnland, D. Repeta, N. Welschmeyer, Improved HPLC method for the analysis of chlorophylls and carotenoids from marine phytoplankton, Mar. Ecol. Prog. Ser. 77 (1991) 183–196. [33] R.L. Airs, J.L. Garrido, Liquid chromatography-mass spectrometry for pigment analysis, in: S. Roy, C.A. Llewellyn, E.S. Egeland, G. Johnsen (Eds.), Phytoplankton Pigments: Characterization, Chemotaxonomy and Applications in Oceanography,

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