Chlorophylls and β‐carotene in Spirulina platensis Dietary Supplements: Nano Liquid Chromatography Versus Diffuse Reflectance Spectroscopy for Establishing Composition, Quality Control, and Comparison of Commercial Products

ABSTRACT

Spirulina is considered a superfood due to its chlorophylls. Two new methods for the determination of chlorophylls and β‐carotene were developed here, one based on in‐tube solid‐phase microextraction (IT‐SPME) coupled online to nanoliquid chromatography (nanoLC) with diode array detection (DAD), and the other on ultraviolet‐visible diffuse reflectance spectroscopy (UV‐vis DRS). A protocol to extract the pigments from spirulina was proposed using ethanol (1.5 mL). The aim is to provide fast and environmentally friendly techniques for both the extraction and measurement of pigments, generating little waste and low energy expenditure. IT‐SPME‐nanoLC‐DAD and UV‐vis DRS showed good linearity up to 300 or 600 µg/L and 50 mg/L, with instrumental detection limits between 10 and 50 µg/L and 2 and 5 mg/L, respectively. The % intraday and interday relative standard deviation were between 2 and 9. The HEXAGON tool is used for assessing greenness and sustainability for three scenarios: establishing composition, quality control, and comparison of seven dietary supplements

1. Introduction

Within the wide range of dietary supplements available in the market, Spirulina (Arthrospira maxima and platensis) is a cyanobacterial microalgae and has become increasingly popular. Among all the active compounds of Spirulina [1, 2], chlorophylls generate the greatest interest for studying, due to their potential antioxidant properties [3]. Microalgae have received great attention because of their capacity to produce photosynthetic pigments with higher efficiency than terrestrial plants [4, 5]. Also, they can be produced on a large scale [4]. In addition to chlorophylls and carotenoids, Spirulina contains other bioactive compounds such as phenolic compounds, essential fatty acids, vitamins, minerals, and highly digestible proteins, all of which contribute to its well‐known nutritional and functional value.

Due to the growing interest in supplementary food, there is a need to develop new analytical techniques that enable the characterization and determination of the active compounds of Spirulina through sustainable, green, and rapid methods. The most used techniques for characterization and determination of chlorophylls in Spirulina are: spectrophotometric methods (ultraviolet‐visible [UV‐vis]) [6, 7, 8, 9, 10], liquid chromatography (LC) with diode array (DAD), mass spectrometry (MS) or UV detection [3, 7, 8, 11, 12, 13, 14, 15, 16, 17], and thin‐layer chromatography (TLC) [6, 15]. Different extraction procedures of chlorophylls from Spirulina were published in the literature [6, 7, 9, 11, 14, 15, 16, 17] from different amounts of samples and solvents, times, and techniques. Here, a comparison of five extraction procedures for chlorophylls and β‐carotene from Spirulina dietary supplements was evaluated from their efficiency in terms of minimizing both extracted mass and extraction time.

In addition, the utilization of miniaturized analysis techniques, like nano LC, has emerged in recent years as an alternative to conventional LC. Nano LC offers various advantages compared to conventional LC, such as reduction in column diameters and particle size, and workflow streamlining, which in turn results in better resolution and sensitivity, shorter times of analysis, reduced solvent consumption, and lower energy consumption [18, 19, 20, 21]. It is also possible to integrate this technique easily with in‐tube solid‐phase micro or nano extraction (IT‐SPM(N)E) [21, 22, 23, 24, 25, 26, 27], named in function of the coating thickness of the capillary, µm or nm, respectively. Due to a considerable number of IT‐SPME applications using both dimensions, and the term nanoextraction was not used, in this paper, IT‐SPME was also employed. This coupling allows the sample volume processed in miniaturized LC to be expanded without compromising resolution. This integration offers additional benefits, such as a reduction in waste generation and increased sensitivity. This makes nano LC an excellent alternative compared to conventional LC [21, 23, 24, 28].

Our research group proposed the determination of chlorophylls in environmental water matrices using IT‐SPME coupled to capillary LC (cap‐LC) and nano‐LC, both equipped with DAD [29, 30]. Here, we demonstrated that the coupling of IT‐SPME‐nanoLC‐DAD is useful to discriminate several pigments in spirulina, both Arthrospira maxima and platensis.

On the other hand, a fast and environmentally friendly technique as UV‐vis diffuse reflectance spectroscopy (DRS), is presented in this work. Some authors have determined chlorophyll through this technique in different matrices, mainly leaves [31, 32, 33]; however, no further information has been found in the literature regarding the use of this technique in Spirulina samples as food supplements.

Figures of merit of both methods are established, and the procedures were compared by using the HEXAGON tool [34, 35]. This tool uses objective criteria divided into five different blocks, namely, figures of merit, toxicity and safety, residues, carbon footprint, and economic cost. For each block, the overall qualification is scaled from 0 to 4, and it is depicted on a regular hexagonal pictogram that allows a user‐friendly comparison of analytical procedures [34]. Eventually, the arithmetic mean (Sav) of the 0–4 scale is computed in order to compare analytical methods from a single data set [35]. HEXAGON is in line with green and sustainable chemistry philosophy, balancing also the figures of merit needed for solving a given problem. The lower the score, the better the adaptation of the analytical procedure to greenness and sustainability aspects that lead to a reliable analytical result.

From the discussion, both IT‐SPME‐nanoLC‐DAD and UV‐vis DRS methods have been qualified for solving different problems to test attending the analytical results that each one can provide for three scenarios: establishing the composition of chlorophylls and β‐carotene, quality control, and comparison of commercial products. The strategies are: miniaturization, minimal consumption of solvents, and selecting, in function of the scenario, the greener and more sustainable option. The present work advances the state of the art in comparison with other procedures in terms of performance, applicability to real‐world samples, wearing in mind the scenario to be solved, sustainability, and greenness.

2. Material and Methods

2.1. Chemicals and Solutions

All reagents used were of analytical grade. Chlorophyll a (95% purity), chlorophyll b (95% purity), and β‐carotene (95% purity) were obtained from Sigma‐Aldrich (St. Louis, MO, USA). HPLC‐grade ethanol was purchased from VWR Chemicals (Radnor, PA, USA).

Stock solutions of the analytes (1000 µg/mL) were prepared by dissolving the appropriate amounts of the commercial standards in ethanol. Working solutions of the analytes and their mixtures were prepared by diluting the stock solutions with ethanol or water; in this last case, the percentage of ethanol in the solutions used for measurement was 20% (v/v). Ultrapure water was obtained from an Adrona system (Riga, Latvia). All the solvents were filtered through 0.22 µm × 47 mm nylon membranes purchased from GVS (Sanford, ME, USA) before use. All solutions were stored in the dark at 4°C until use.

2.2. Chromatographic Conditions

The NanoLC system was an Agilent 1260 Infinity (Waldbronn, Germany) equipped with an injection system consisting of a Rheodyne 7725i 6‐position, 1/16″ manual injection valve connected to a VICI C2N 10‐port 2‐position, 1/32″ automatic valve (Valco Instruments, Houston, TX, USA) and a UV‐vis DAD detector (Agilent). The standard solutions were preconcentrated online through in‐valve IT‐SPME using a fused silica capillary (10 cm length × 75 µm i.d.) (Análisis Vínicos, Tomelloso, Spain) coated with a polymer of tetraethyl orthosilicate (TEOS), triethoxymethylsilane (MTEOS) doped with SiO₂ nanoparticles. A homogeneous coating (350 nm) of TEOS‐MTEOS was formed inside the capillary column [27]. The IT‐SPME was coupled to the analytical column (Zorbax 300SB‐C18 5 cm × 75 µm i.d., 3.5 µm particle size) from Agilent using the setup given in Figure 1. The procedure begins with standard or sample loading from the manual valve (V1) to the automatic valve (V2), which contains the IT‐SPME capillary. During the preconcentration stage, the analytes are retained on the capillary coating, while the remaining solution is directed to waste prior to chromatographic separation. In addition, the analytical column is conditioned (position A of Figure 1). After, in position B, the crude IT‐SPME extract was injected from the automatic valve V2 in the chromatographic system, and the registers were obtained.

FIGURE 1.

Schematic of the injection system for the in‐tube solid‐phase microextraction (IT‐SPME)‐nano‐LC. V1 and V2 are manual 6‐port and 2‐position, and automatic nano 10‐port and 2‐position injection valves, respectively. In position A, the standard or sample is loaded manually from V1 to V2, and the analytical column is conditioned, and in position B, the crude IT‐SPME extract is injected into the chromatographic system. Dimensions (i.d., length, and volume) for the different parts are: 1) IT‐SPME capillary (75 µm, 10 cm, and 433 nL), 2) connection between V2 and Analytical column (25 µm, 10 cm, and 49 nL), 3) analytical column C18 (75 µm, 5 cm, and 133 nL), and 4) connection between analytical column and UV‐vis DAD (25 µm, 20 cm, and 98 nL). Total volume system 713 nL.

The chromatographic separations were carried out in isocratic elution mode with a mixture of ethanol‐ultrapure water (EtOH/H₂O 95:5% v/v) as mobile phase at a flow rate of 0.5 µL/min. Aliquots of 100 µL of the working solution were manually introduced into the IT‐SPME capillary system using a 250 µL precision syringe from V1 (Figure 1). Once the sample was loaded into the system, the automatic valve was rotated, automatically changing its position, causing the retained analyte to be eluted to the analytical column by the mobile phase.

The analytical signal was recorded between 400 and 800 nm, and monitored at 662 and 420 nm for chlorophyll a, 662 and 450 nm for chlorophyll b, and at 450 nm for β‐carotene. The detector was coupled to a data system (Agilent, ChemStation) for data acquisition and treatment. All the experiments were carried out at room temperature, 20 ± 2°C, throughout the study.

2.3. Spectroscopic Conditions

UV‐vis spectra in reflectance mode were registered with a Cary 60 UV‐vis spectrophotometer (Agilent Technologies, Santa Clara, CA, USA) equipped with a diffuse reflectance probe from Harric Scientific Products. Spectra were recorded from 200 to 800 nm, and monitored at 662 nm and 420 nm for chlorophyll a, 662 and 450 nm for chlorophyll b, and at 350 and 450 nm for β‐carotene. For data collection and processing, CaryWinUV software was used. The measurement involved directly applying 50 µL of either a standard ethanolic solution or Spirulina platensis ethanolic extract on a Millipore fibre‐glass filter 13 × 1.2 mm (Sigma‐Aldrich, Darmstadt, Germany). This was carried out in low‐light settings, with immediate measurements taken directly on the filter.

2.4. Extraction Procedure

Five protocols (A, B, C, D, and E) of sequential ethanol extractions of the pigments from dietary supplements (1 mg and 10 mg) as described in Table 1 were carried out. The lixiviation was aided by vortex agitation for 1 min, followed by 5 min in an ultrasonic bath. The obtained supernatants were filtered with 0.2 µm × 13 mm nylon membrane filters and diluted with water, ensuring that the final ethanol content in each solution was adjusted to 20% v/v and processed by IT‐SPME‐nanoLC‐DAD. If necessary, the ethanol volume was adjusted accordingly to maintain this ratio.

TABLE 1.

Tested protocols for the extraction of pigments from dietary supplements.

Extraction	Sample mass, mg	Number of extractions × volume (mL)
A	1	2 × 0.2 1 × 1.2 3 × 0.2
B	1	1 × 0.2 8 × 0.15
C	1	9 × 0.5
D	10	9 × 0.5
E	1	1 × 0.2 1 × 1.3 3 × 0.2

Amber material was used to prevent light exposure. Samples and standards were prepared on the same day and injected immediately after the extraction. Table 1 shows the different protocols for the extraction of pigments from dietary supplements.

2.5. Samples of Spirulina

Seven Spirulina products purchased from various local markets were analysed. Each product came with specific details listed on its label, as summarized in Table 2. This table also shows the weighted amount for extraction procedure E (see Table 2) and the dilution factor employed for IT‐SPME‐nanoLC‐DAD. Three replicates were measured from the extract directly in a fibre‐glass filter for UV‐vis DRS and diluted with ultrapure water for IT‐SPME‐nanoLC‐DAD and maintaining a content of ethanol of 20%. Table 3 indicates the kinds of microalgae. Product S1 was presented as powder, and the other ones as capsules.

TABLE 2.

Characteristics and nutritional content of the Spirulina samples analyzed.

Sample/Weighted amount (mg)	Microalgae	Net weight (mg)	Spirulina (mg)	Nutritional composition label per 100 g	Dilution factor NanoLC
S1/1.30	Spirulina (Arthrospira maxima)	300 (per dosis)	300 (per dosis)	Fat 3.1 g, saturated fat 0.65 g, carbohydrates 12 g, sugar 1.4 g, fibre 9.2 g, protein 12 g, and salt 0.01g	10
S2/0.91	Spirulina (Arthrospira platensis)	300 (per capsule)	296 (per capsule) 7.90 mg g−1 a	Fat 6 g, carbohydrates 15.1 g, sugar 0.5 g, fibre 6.47 g, protein 62.5 g, and salt 2.7 g.	19
S3/1.22	Spirulina (Arthrospira maxima)	458 (per capsule)	350 (per capsule)	Fat 2.7 g, saturated fat 1.4 g, carbohydrates <0.5 g, sugar <0.5 g, fibre 11 g, protein 71 g, and salt 1.7 g	10
S4/1.22	Spirulina (Arthrospira maxima)	400 (per capsule)	357 (per capsule)	Fat 5 g, carbohydrates 5 g, and protein 60 g	10
S5/0.99	Spirulina (Arthrospira platensis)	510 (per capsule)	410 (per capsule)	Fat 8 g, carbohydrates 24 g, and protein 57 g	10
S6/1.15	Spirulina (Arthrospira platensis)	570 (per capsule)	300 (per capsule)	Total fats 8 g, carbohydrates 24 g, protein 60%–70%, vitamin A, and vitamin C	10
S7/1.83	Spirulina (Arthrospira platensis)	400 (per capsule)	312 (per capsule)	Protein 65–70%, vitamin E, and B complex vitamins	19

^a Chlorophyll a.

TABLE 3.

Instrumental analytical parameters of the in‐tube solid‐phase microextraction (IT‐SPME)‐nanoLC‐DAD.

Linearity, y = b1 x + b0 (n = 10)
Compound	Interval (mg/L)	b1 ± sb1(L/mg)	b0 ± sb0	R2	LOD (mg/L)	RSD% (n = 3)*
Chlorophyll a	0.03–0.3	(2190 ± 40)a	−11 ± 5	0.998	0.01	2d/5e
	0.03–0.3	(2051 ± 40)b	−29 ± 5	0.998	0.01	2d/5e
Chlorophyll b	0.06–0.6	(1098 ± 40)c	31 ± 11	0.992	0.02	3d/7e
	0.15–0.6	(325 ± 11)b	32 ± 6	0.991	0.05	3d/7e
β‐carotene	0.03–0.3	(2595 ± 50)c	2 ± 8	0.997	0.01	3d/8

*Concentration of 100 µg/L for chlorophyll a, 300 µg/L for chlorophyll b and 200 µg/L for β‐carotene and intraday RSD%. Established at: ^a420 nm, ^b 662 nm, and ^c 450 nm. ^dIntraday precision and ^eInterday precision.

To minimize degradation from light exposure, each sample was stored in amber Eppendorf tubes. These samples were then kept in a dark environment at room temperature to preserve their integrity and ensure the accuracy of the analysis.

Matrix effect was also evaluated from the first extracts of Spirulina samples, which contained a minimal amount of analytes. A standard solution with a concentration of 50 µg/L for chlorophyll a and β‐carotene, and 100 µg/L for chlorophyll b was added to these samples.

3. Results and Discussion

3.1. Figures of Merit of IT‐SPME Coupled On‐line With nanoLC‐DAD

The nanoLC system shown in Figure 1 was used for all experiments. Figure 2 gives the chromatogram for standards of chlorophylls b and a and β‐carotene at 662 and 450 nm. The inserts correspond to the normalized spectra obtained at the maximum of each chromatographic peak for the pigments. The run was less than 8 min at a flow rate of 0.5 µL/min.

FIGURE 2.

Chromatogram of standard solutions of (A) chlorophylls b and a (300 µg/L measured at 662 nm) and (B) chlorophyll b 300 µg/L and β‐ carotene 200 µg/L measured at 450 nm. The inserts show the respective normalized UV‐vis spectra at the maximum of the corresponding chromatographic peak.

The calibration curves were obtained using standard solutions of the analytes. For processing samples, the pigment recoveries from the extraction protocol applied to the supplements should be considered (this information appears later in the paper). Instrumental linearity was evaluated by processing in duplicate five concentrations of chlorophyll a, chlorophyll b, and β‐carotene prepared with a 20% ethanol solution. The two values for each concentration were statistically similar (y = x, R² = 0.99). It can be observed that a good correlation between peak areas and the concentration of the different pigments is obtained for the external calibration, which is reflected in the R² value for all calibration lines (Table 3). Sufficient evidence of selectivity and the absence of significant matrix effects will be evaluated from peak purity analysis in the samples section, thereby supporting the use of external calibration with standard solutions if values near 1000 are obtained.

Detection limits (LODs) and quantification (LOQs) were obtained by gradually injecting lower concentrations of diluted standard until achieving a signal‐to‐noise ratio of 3 and 10, respectively, always maintaining 20% of ethanol. These values are given in Table 3; the LOQ was the lower value of the working interval of concentrations. The intraday precision (% relative standard deviation [%RSD]) was also suitable, being between 2% and 3%. The interday precision achieved was between 5% and 8%.

3.2. Figures of Merit of UV‐vis Reflectance Spectroscopy

Chlorophyll a, chlorophyll b, and β‐carotene standards in ethanol were used to construct calibration curves using their spectra. Two absorption maxima were observed for each of the chlorophyll standards. Chlorophyll a exhibited peaks at approximately 430 and 660 nm, while chlorophyll b showed maxima around 470 and 650 nm. For β‐carotene, the absorption maximum occurred around 350 nm. The spectra obtained are shown in Figure 3.

FIGURE 3.

Spectra of ethanolic standards of chlorophyll b (Chlo.b), and chlorophyll a (Chlo.a) at 35 mg/L and β‐carotene at 50 mg/L.

For diffuse reflectance, linearity was evaluated by processing in duplicate three concentrations of chlorophyll a, chlorophyll b, and β‐carotene (Table 4). These values were obtained by measuring directly the standard solution on fiberglass filters (see inserts of Figure 3). For processing samples, the pigment recoveries from the extraction protocol applied to the supplements should be considered (this information appears later in the paper). The LODs and LOQs were obtained from the standard deviation of the signals of ten blanks (see Table 4); the LOQ was the lower value of concentration of the working interval of concentrations. The intraday precision (RSD%) was between 2% and 6% and the interday precision (RSD%) was between 5% and 9%. These RSD% values are suitable.

TABLE 4.

Instrumental analytical parameters of the diffuse reflectance spectroscopy.

Linearity, y = b1 x + b0 (n = 6)
Compound	Interval (mg/L)	b1 ± sb1 (L/mg)	b0 ± sb0	R2	LOD (mg/L)	RSD% (n = 3)*
Chlorophyll a	6–50	(0.0045 ± 0.0003)a	0.008 ± 0.006	0.99	2	5 g/9h
	6–50	(0.0037 ± 0.0002)b	0.05 ± 0.01	0.985	2	3 g/7h
Chlorophyll b	15–50	(0.0040 ± 0.0002)c	0.09 ± 0.001	0.990	5	6 g/8h
	15–50	(0.0013 ± 0.0001)d	0.06 ± 0.003	0.99	5	2 g/5h
β‐carotene	6–50	(0.0132 ± 0.0002)e	0.01 ± 0.02	0.995	2	5 g/8h
	6–50	(0.0154 ± 0.0005)f	0.01 ± 0.03	0.990	2	2 g/6h

*Concentration of 35 mg/L for chlorophyll a, 25 mg/L for chlorophyll b and for β‐carotene. Established at: ^a432 nm, ^b667 nm, ^c472 nm, ^d653 nm, ^e450 nm, and ^f350 nm. ^gIntraday precision and ^hInterday precision.

3.3. Study of the Extraction Procedure

The extraction protocols given in Table 1 were assayed for sample S1, and chlorophyll a was selected because of its higher presence in the samples. Successive extractions with ethanol of S. platensis were tested until the extract coloration became colourless. The presence of chlorophyll a was confirmed in each of the extracts using the IT‐SPME‐nanoLC‐DAD technique until they were below the LOD. Five protocols were established (A, B, C, D, and E), and the strategy was to minimize the number of extractions and sample and ethanol amounts. A and B used a total volume of ethanol of 2.2 and 1.4 mL, respectively, and tested the influence of the volume of the several extractions on the pigment recovery. C and D evaluated the influence of sample amount and maintained el number of extractions and their volume, and the total volume of ethanol was 4.5 mL. E is similar to A, but the number of extractions decreases. Figure 4 shows the amount of recovered chlorophyll a in each of the several extraction procedures (A, B, C, D, and E), with the X‐axis representing each extraction step and the Y‐axis indicating the amount of chlorophyll a recovered. Figure 5 gives the total amount recovered of chlorophyll a for each protocol. A and E provided higher recoveries of the pigment for the first two steps than B, C, and D.

FIGURE 4.

Number of extractions vs recovered amount (mg) of chlorophyll a for sample S1.

FIGURE 5.

Representation of the chlorophyll a quantities (mg/g) and the total volume of ethanol (EtOH) in mL for sample S1, obtained across the different extractions performed.

The accumulative values obtained in extraction A, B, C, D, and E for chlorophyll a in sample S1 were: 4.01 ± 0.92, 4.1 ± 0.2, 4.6 ± 0.2, 4.7 ± 0.4, and 4.7 ± 0.2 mg/g, respectively (Figure 5). It can be observed that across all extractions, the quantities of analytes obtained were relatively similar. For extractions C and D, a significantly larger volume of solvent (4.5 mL of ethanol) and a higher number of extraction cycles (nine in total) were used compared to extractions A, B, and E, which utilized 2.2, 1.4, and 2.1 mL of ethanol, respectively. However, extraction E stands out as it required only 2 extraction cycles to achieve comparable results, suggesting that a more efficient extraction process can be achieved with fewer steps and less solvent. We selected protocol E with the first two extractions (1.5 mL) for processing samples.

Table 5 shows different extraction procedures of chlorophylls from Spirulina published in the literature [6, 7, 9, 11, 14, 15, 16, 17]. As can be seen very different number of samples and solvents, times, and techniques were proposed. Here extraction method was developed with a minimal use of a friendly solvent as ethanol, and using around 1 mg of sample.

TABLE 5.

Published extraction procedures for chlorophylls from Spirulina.

Reference	Kind of sample	Extraction time (min)	Sample mass	Solvent	Method of extraction
[6]	Commercial dietary supplements	20	100 mg sample + 10 mg MgCO3	1 mL acetone	Ultrasonic bath, microtube homogenizer, centrifuge
[7]	Spirulina (Arthrospira maxima)	40	0.025 g	1 mL EtOH	Stirring (Orbital mixer) and centrifuge
[9]	Spirulina	1, 2, or 3	0.5 g	35 mL EtOH	Three different ultrasonic extractions
[11]	Algal pellets	Overnight	0.4 or 0.9 g	1 mL acetone, 10% (w/v) *TCA, 0.07% *DTT	Centrifuge
[14]	Algae Chlorella vulgaris	40	100 mg	50 mL EtOH	Vortex, Ultrasonic bath, and rotary evaporator
[15]	Spirulina (Arthrospira platensis)	3, 9, or 15	2.5 g of sample	11 mL of EtOH, Hexane, or Petroleum ether	*ASE
[16]	Spirulina (Arthrospira platensis)	No information	75 g	CO2 with 10% EtOH	*SFE
[17]	Spirulina (Arthrospira platensis)	15	10 g	100 mL EtOH	Ultrasonic cell and centrifuge
This Work	Spirulina (Arthrospira platensis or maxima)	6	1 mg	1.5 mL EtOH	Vortex and ultrasonic bath

*TCA = trichloroacetic acid; *DDT = dithiothreitol; *ASE = assisted solid extraction; *SFE = Supercritical fluid equipment.

3.4. S. platensis Dietary Supplements: Pigment Composition, Quality Control, and Product Comparison

Matrix effect was evaluated by IT‐SPME‐nanoLC‐DAD from the first extraction of Spirulina samples, which contained a minimal amount of analytes, which were spiked with 50 µg/L for chlorophyll a and β‐carotene, and 100 µg/L for chlorophyll b. The results obtained (Table 6) indicate the absence of matrix effect, suggesting that calibration can be performed externally without compromising the accuracy of the estimated concentrations.

TABLE 6.

Recovery and precision obtained in fortified samples with 50 µg/L of chlorophyll a and β‐carotene and 100 µg/L for chlorophyll b in the first sample extract of Spirulina platensis.

Sample	Recovery % (n = 3)
	Chlorophyll a	Chlorophyll b	β‐carotene
S1	85 ± 8	89 ± 12	93 ± 1
S2	108 ± 12	85 ± 10	91 ± 3
S3	99 ± 9	85 ± 10	91 ± 1
S4	102 ± 5	91 ± 7	86.1 ± 0.1
S5	97 ± 8	88 ± 9	96 ± 6
S6	91 ± 2	82 ± 5	94 ± 2
S7	110 ± 5	87 ± 10	88.8 ± 0.2

Figure 6 shows the chromatograms of the second extract of all samples (S1–S7) processed using the extraction procedure E, which gives the maximum extraction of the pigments (see Figure 5) and is diluted properly. The peak purities were for chlorophyll A: 979 (S1), 963 (S2), 961 (S3), 987 (S4), 978 (S5), 978 (S6), and 926 (S7), and for β‐carotene: 989 (S3) and 971 (S4). These values indicate that external calibration based on the selectivity of the DAD detector provides suitable results.

FIGURE 6.

Chromatograms obtained at 662 nm (blue line) and 450 nm (red line) for real samples: S1, S2, S3, S4, S5, S6, and S7. The chromatogram shows the following peaks: 1) chlorophyll b, 2) chlorophyll a, 3) β‐carotene, and 4*) an additional compound reported in the literature as a chlorophyll derivative found in microalgae [1].

The average value of peak widths at 4σ (ω) obtained by the software of the nanoLC instrument was: 0.39 ± 0.06 min (n = 9), which is in accordance with an inherent trade‐off in the system design (see Figure 2), which prioritizes enhanced detection limits and minimal solvent consumption, in accordance with a green analytical approach. The coupling IT‐SPME‐nanoLC permits to injection into the analytical column a volume of 433 nL of processed sample or standard, being the total volume of the system 713 nL as indicated in Figure 2 with suitable resolution.

The most predominant component in the tested samples was chlorophyll a, identified by its spectrum and retention time in accordance with Figure 2, which is consistent with the literature, as chlorophyll a is the primary pigment found in S. platensis. In addition, samples S1, S2, S3, and S4 also contained a compound reported in the literature as chlorophyll in microalgae [1]. The insert of Figure 6 gives its spectrum obtained at the maximum of the chromatographic peak and is named as unknown chlorophyll, which is similar to that given in [1]. Samples S5, S6, and S7 did not exhibit the presence of this compound. β‐carotene was detected in samples S3 and S4; however, in sample S3, its concentration was near LOQ, while in sample S4 it was higher. Additionally, samples S3–S7 displayed the presence of other carotenoids, as indicated by their characteristic spectral signatures. However, these compounds could not be definitively identified due to the lack of analytical standards. Despite the need for prior dilution and the fact that this is not a direct measurement technique, the high sensitivity and ability to identify multiple compounds of interest make it an ideal method for the composition studies. Table 7 shows the pigment composition estimated for all samples. The amount of chlorophyll a found in the samples was between 4.7 ± 0.2 and 7.1 ± 0.3 mg/g, similar to that reported in the literature and to that found using the IT‐SPME technique proposed in this study [6, 7, 9, 11, 12].

TABLE 7.

Concentrations obtained from chlorophyll a (Chlo.a), chlorophyll b (Chlo.b), Chl. from microalgae, and β‐carotene in in‐tube solid‐phase microextraction (IT‐SPME) coupled to nanoLC and diffuse reflectance for the various Spirulina samples. All chls. are estimated as chlorophyll a by diffuse reflectance.

Concentration mg/g IT‐SPNE coupled to nanoLC‐DAD						Concentration mg/g Diffuse reflectance
Sample	Chlo.a	Chlo.b	Chl. from microalgae	β‐carotene	Other carotenes	Chls. at 667 nm	Carotenes
S1	4.7 ± 0.2	ND	Detected	ND	ND	5.8 ± 0.9	ND
S2	6.8 ± 0.3	ND	Detected	ND	ND	7.4 ± 0.8	ND
S3	1.6 ± 0.1	ND	Detected	0.03 ± 0.01	Detected	<2	Detected
S4	4.3 ± 0.7	ND	Detected	0.31 ± 0.05	Detected	5.8 ± 0.3	Detected
S5	5.9 ± 0.3	ND	ND	ND	Detected	6.0 ± 0.4	Detected
S6	4.7 ± 0.4	ND	ND	ND	Detected	6.2 ± 0.2	Detected
S7	7.1 ± 0.3	ND	ND	ND	Detected	7.7 ± 0.3	Detected

*ND: non‐detected.

Figure 7 gives the reflectance diffuse spectra for the assayed samples. The spectra of samples S1, S2, and S3 are in accordance with the chlorophyll a spectrum (Figure 3). Those results are in accordance with the chromatograms of Figure 6, S1 and S2 only contain chlorophylls, and S3 contains a low concentration of β‐carotene too (see Table 7). The spectra of S4, S5, S6, and S7 show a mixture of spectra of chlorophylls and carotenoids, which are also in accordance with their chromatograms.

FIGURE 7.

Spectra of samples S1, S2, S3, S4, S5, S6 and S7 obtained by diffuse reflectance spectroscopy.

Diffuse reflectance spectroscopy can give total concentrations of pigments expressed as chlorophyll a here. The results obtained are shown in Table 7.

IT‐SPME‐nanoLC‐DAD and UV‐vis DRS methods have been qualified for solving different problems to test attending the analytical results that each one can provide: establishing the composition of chlorophylls and β‐carotene, quality control, and comparison of commercial products. Figure 8 indicates that IT‐SPME‐nanoLC‐DAD can be employed for the three problems established. However, considering the HEXAGON tool [34, 35] obtained for the LC method and that corresponding to the UV‐vis DRS, this second method is more sustainable and greener. HEXAGON, as mentioned in the introduction section, is in line with the green and sustainable chemistry philosophy, also balancing the figures of merit needed for solving a given problem. Objective criteria are evaluated through the definition of penalty points (PPs) divided into five different blocks, namely, figures of merit, toxicity and safety, residues, carbon footprint, and economic cost. For each block, the overall qualification is scaled from 0 to 4, and it is depicted on a regular hexagonal pictogram that allows a user‐friendly comparison of analytical procedures [34, 35]. Figure 9 shows the PPs for each method for the several variables employed by the HEXAGON tool. The UV‐vis DRS method showed lower penalties in most of the analyzed categories, particularly in terms of toxicity, waste generation, carbon footprint [36, 37], and annual cost, highlighting its environmentally friendly character. References 34 and 35 give a detailed explanation of how to calculate PPs. Therefore, although the IT‐SPME‐nanoLC‐DAD method provides a higher resolving power for compositional analysis of complex matrices like spiruline, the UV‐vis DRS method represents a greener alternative for specific purposes such as routine quality control and product comparison, as also shown in Figure 8. This allows for the selection of the most appropriate method not only based on analytical performance but also considering environmental impact and practical feasibility.

FIGURE 8.

Utilities of in‐tube solid‐phase microextraction (IT‐SPME)‐nanoLC‐DAD and ultraviolet‐visible diffuse reflectance spectroscopy (UV‐vis DRS) for contributing to evaluating pigment composition, quality control, and comparison of the market products. HEXAGON tool for each method. See text for more explanations.

FIGURE 9.

Variables considered for establishing the HEXAGON tool for in‐tube solid‐phase microextraction (IT‐SPME)‐nanoLC‐DAD and ultraviolet‐visible diffuse reflectance spectroscopy (UV‐vis DRS).

4. Conclusions

The extraction of chlorophylls and β‐carotene in S. platensis is quite efficient compared to others reported in the literature. It stands out for the small amount of sample required, the minimal solvent used, the short time employed, and the number of extractions needed to extract chlorophyll in the samples.

In this study, the determination of chlorophylls in S. platensis was assessed by means of IT‐SPME coupled to nanoLC‐DAD and UV‐vis DRS. IT‐SPME‐nanoLC‐DAD provided a quick analysis of chlorophylls and β‐carotene (less than 8 min), low energy cost, and minimal solvent usage. The methodology demonstrated strong linearity and accuracy, with satisfactory intraday and interday precision. It's noteworthy for evidence by minimal solvent usage in mobile phase utilization, along with swift analysis time in reference to conventional LC. IT‐SPME‐nanoLC‐DAD offers high sensitivity and the possibility to detect multiple analytes. Diffuse reflectance it's a quick and efficient technique. Its sustainability and eco‐friendly usage, and ease of use are also worth mentioning, as qualified by the HEXAGON tool. This technique offers a comprehensive approach to the characterization of complex mixtures, making them invaluable for the detailed analysis of multicomponent samples. The LC method is useful for giving a response for establishing the composition of chlorophylls and β‐carotene, quality control, and comparison of commercial products. Meanwhile, UV‐vis DRS can be employed for quality control and comparison of products.

Author Contributions

C. Soto: investigation, data curation, methodology, and writing–original draft; R. Herráez‐Hernández: methodology, data curation, and writing–review & editing;  P. Campíns‐Falcó: data curation, methodology, supervision, writing–original draft, funding acquisition, writing—review & editing, and project administration.

Conflicts of Interest

The authors declare no conflicts of interest.

Soto C., Herráez‐Hernández R., and Campíns‐Falcó P., “Chlorophylls and β‐carotene in Spirulina platensis Dietary Supplements: Nano Liquid Chromatography Versus Diffuse Reflectance Spectroscopy for Establishing Composition, Quality Control, and Comparison of Commercial Products.” Journal of Separation Science 48, no. 9 (2025): 48, e70260. 10.1002/jssc.70260