Evaluation of the Effect of Growth Factors on Chlorophylls a and b Production from Microalgae

Summary

The present study highlights the advantages of employing the method developed by Jeffrey and Humphrey for extracting and quantifying fat-soluble pigments from microalgae. This method serves as a valuable tool for assessing the influence of growth factors on chlorophyll production and cellular content in these organisms.

Abstract

Microalgae contain two main groups of pigments: chlorophylls and carotenoids. Chlorophyll is a green pigment that absorbs light energy and transforms it into chemical energy to facilitate the synthesis of organic compounds. This pigment serves as a valuable primary source for biotechnological input products in the food, pharmaceutical, and cosmetic industries due to its high antioxidant properties and coloring capabilities. The objective of this research was to evaluate the effect of growth factors (CO₂ concentration, light color, and light intensity) through a Taguchi L₄ experimental design on cell growth and the cellular content of chlorophyll a and b in Chlorella sorokiniana, followed by validation of the method using Haematococcus pluvialis microalgae as an additional study model. Cell growth was quantified using the optical density spectrophotometric technique at a wavelength of 550 nm. For the quantification of chlorophylls, a cell extract was obtained using a 90% pure acetone solution, and subsequently, the concentrations of chlorophylls a and b were quantified using spectrophotometric techniques at wavelengths of 647 nm and 664 nm, according to the method described by Jeffrey and Humphrey. The experimental results indicated that controlling conditions of low CO₂ addition, purple light, and low light intensity increases both cell growth and the concentration of chlorophylls a and b within the cells. The implementation of this chlorophyll quantification method allows for quick, simple, and precise determination of chlorophyll content, as the wavelengths used are at the absorbance peaks of both types of chlorophylls, making this technique easily reproducible for any microalgae under study.

Introduction

In recent years, the growing environmental problems caused by anthropogenic activities and their adverse effects on the health and balance of ecosystems have driven the search for more efficient and environmentally friendly production systems. This has accelerated processes in industries and fostered the implementation of bioremediation treatments and the development of bio compounds to mitigate these harmful effects¹.

This context has led to a significant growth in the study of microalgae, driven by the need to find innovative solutions to current environmental and economic challenges. Microalgae thrive in aquatic environments, using sunlight and carbon dioxide as sources of energy and carbon, respectively. This characteristic makes them a sustainable and promising alternative for producing various valuable products. Research in this field has focused on understanding the physiology and metabolism of these cells, as well as developing efficient technologies for their cultivation and processing².

There is a clear need to develop accessible and reliable tools for studying microalgae to accelerate research processes and deepen the understanding of their physiology, metabolism, and potential applications. These tools should enable rapid analysis of the effect of environmental and production factors on key parameters, such as chlorophyll concentration, which is a fundamental indicator of the health and development of these organisms. A decrease in chlorophyll content may indicate environmental stress, nutritional deficiencies, or diseases.

These green pigments play a crucial role in photosynthesis, capturing sunlight energy to convert carbon dioxide and water into glucose and oxygen and constitute between 0.5% and 5% of the biomass of microalgae³. Beyond their essential role in sustaining life processes, chlorophylls have found applications in various industries. Chlorophyll extracts are utilized in food and beverage processing as natural colorants, imparting vibrant green hues to products while also providing antioxidant properties. Moreover, chlorophyll-based supplements are gaining popularity in the health and wellness sector due to their purported detoxifying and anti-inflammatory effects. By harnessing the multifaceted properties of chlorophyll, industries can develop innovative products that contribute to both visual appeal and consumer well-being¹.

In this regard, one microalgae of great biotechnological relevance is C. sorokiniana. This microorganism stands out for its rapid growth rate, making it highly efficient in biomass production. Additionally, C. sorokiniana exhibits a diverse content of highly nutritional compounds, including proteins, lipids, and vitamins, which renders it valuable for various applications in food, feed, and biofuel production². Moreover, this microalgae species has been found to produce extracellular enzymes with diverse functions, opening up possibilities for biotechnological applications such as wastewater treatment, bioremediation, and pharmaceuticals⁴. In addition to its rapid growth and versatile applications, C. sorokiniana also demonstrates significant potential for chlorophyll production. As a photosynthetic microorganism, C. sorokiniana possesses the machinery necessary for the synthesis of chlorophyll, which constitutes between 0.5% and 5% of the biomass of microalgae³. This capability makes C. sorokiniana an attractive candidate for commercial-scale chlorophyll production and promises to be a sustainable solution to urgent environmental and nutritional challenges⁵.

On the other hand, another microalgae species of significant interest is H. pluvialis. This microalga is renowned for its production of astaxanthin, a potent antioxidant pigment with numerous industrial applications. Astaxanthin serves as a protective mechanism for H. pluvialis photosystem, shielding it from oxidative stress induced by environmental factors. This pigment is highly sought after in cosmetics, nutraceuticals, and aquaculture industries, owing to its antioxidant properties and potential health benefits. With its abundant ability to produce astaxanthin, H. pluvialis presents a promising avenue for developing innovative products catering to various industrial and consumer needs⁶.

Recent studies have also highlighted H. pluvialis¹ potential for chlorophyll production, further enhancing its industrial significance. For instance, a study investigated the chlorophyll content of H. pluvialis under varying growth conditions and found that under optimal cultivation parameters, H. pluvialis exhibited remarkable chlorophyll production rates, surpassing those of other microalgae species⁷. This finding underscores the potential of H. pluvialis as a sustainable source of chlorophyll, offering novel opportunities for its utilization in various industrial sectors.

Protocol

1. Culture media preparation and inoculum preparation

1. Prepare 1 L of 3N-BBM+ V(CCAP) growth medium (Macronutrients: NaNO₃ [0.75 g L^-1], CaCl₂ [0.019 g L^-1], MgSO₄ [0.019 g L^-1], K₂HPO₄ [0.057 g L^-1], NaCl [0.025 g L^-1], KH₂PO₄ [0.175 g L^-1]; micronutrients: Na₂ EDTA [0.0186 g L^-1], FeCl₃ [0.0024 g L^-1], MnCl₂ [0.001 g L^-1], ZnCl₂ [1.25 x 10^-4 g L^-1], CoCl₂ [5 x 10^-5 g L^-1], Na₂MoO₄ [9.98 x 10^-5 g L^-1])⁸ and sterilize it for 15 min at 121 °C and 1.5 psi.
2. Inoculate a flask with 1 L of growth medium at an approximate concentration of 3 x 10⁶ cells mL^-1 for the microalga C. sorokiniana; or add 5% to 10% (v/v) of inoculum to the growth medium.
   NOTE: To maintain sterile conditions, it is recommended to use a laminar flow hood, gloves, lab coat, and face mask.
3. Incubate the inoculated flask under red light sources with an approximate intensity of 3,600 to 4,200 lux, at a temperature between 22-26 °C, with a photoperiod of 12 h light and 12 h darkness for C. sorokiniana and photoperiod of 16 h light and 8 h darkness for H. pluvialis, and manually agitate every 24 h, approximately for 7 days for both strains.
   NOTE: Maintain under growth conditions for at least 7-10 days (until obtaining an intense green color) for C. sorokiniana and 10-12 days for H. pluvialis. To obtain the microalgae concentrate that will serve as the inoculum in the kinetics, it is necessary to remove the supernatant medium from the produced algae.
4. In a sterile environment, fill sterile 15 mL conical centrifugation tubes with approximately 10 mL of inoculated medium and centrifuge at 3,000 x g for 20 min under 25-30 °C.
5. Under sterile conditions, remove the supernatant using a micropipette or by pouring and concentrating the biomass in a sterile 50 mL conical centrifugation tube.
6. Repeat this process as necessary until the required volume of inoculum for the kinetics is obtained.
7. Measure the optical density (OD₅₅₀ nm) of the inoculum to determine the cell concentration of the microalgae and store it at a temperature of 3-4 °C until ready for use.
   NOTE: Direct cellular counting was initially performed to correlate cell concentration with absorbance. Once counted, the inoculum can be stored under refrigeration for no more than 4 days.

2. Study of the growth kinetics

1. Prepare the required volume of 3N-BBM+ V(CCAP) suitable for each experiment (8 L for photobioreactor level tests for C. sorokiniana and 0.5 L for the flask-level tests for H. pluvialis) according to standard laboratory procedures, ensuring proper sterilization.
2. For both photobioreactor and flasks, configure the light source to the desired color spectrum (Blue at 460 nm, purple, mix of 460 nm and 630 nm for C. sorokiniana and red at 630 nm for H. pluvialis); cover both systems to avoid external light interference.
   NOTE: Due to the variation in wavelength emitted from each light-emitting diode, spectroradiometry was performed using a spectroradiometer on the light sources to determine the wavelength and emission power of each diode⁹. Purple light was achieved by combining red and blue LEDs, resulting in wavelengths of 460 nm with an emission power of 1.65 W nm^-1 for the blue LEDs and 630 nm with an emission power of 0.6 W nm^-1 for the red LEDs. These measurements were used for the tests conducted solely under blue and red light.
3. Program the timer to alternate between 12 h light and 12 h dark cycles for C. sorokiniana and 16 h light and 8 h dark cycles for H. pluvialis.
4. Connect the CO₂ aeration system to the photobioreactor culture vessel to provide carbon dioxide supply in intervals of 12 h for C. sorokiniana.
5. Inoculate the culture vessel with a sufficient volume of pre-inoculum to achieve a starting cell density of 3 x 10⁵ cells mL^-1 or its equivalent in optical density (OD₅₅₀ nm) = 0.180.
   NOTE: Ensure aseptic technique during inoculation to prevent contamination.
6. Sample the culture at regular intervals of 12 h for C. sorokiniana and 8 h for H. pluvialis to monitor cell growth and physiological parameters. Collect samples using sterile techniques to maintain the integrity of the culture.

3. Quantification of chlorophylls

NOTE: Data for cellular quantification is provided in Supplementary File 1.

1. Place 40 mL of the sample into plastic conical centrifugation tubes.
2. Centrifuge at 3,000 x g for 20 min at 15 °C and discard the supernatant by decantation or using a Pasteur pipette.
3. Transfer the cell pellet into a glass tube covered with aluminum foil and a screw cap to prevent oxidation.
4. Resuspend the cell pellet with 3 mL of 90% pure acetone solution using a vortex.
5. Sonicate the suspended sample in an ice bath for two cycles of 5 min each and let it rest at 4 °C for 16 h¹⁰.
6. After 16 h, sonicate again under the same conditions mentioned in the previous step (step 3.5) and centrifuge again under the previously described conditions (step 3.2).
7. Separate the pigment extract using a Pasteur pipette and transfer it to another clean tube protected from light.
8. Place the pigment extract in a quartz cell to read in a spectrophotometer at 630 nm, 647 nm, and 664 nm wavelengths.
   NOTE: Calibrate the spectrophotometer beforehand with 90% acetone.
9. To calculate chlorophyll concentrations, use the following equations¹¹:
   Chlorophyll a = 11.93 A₆₆₄ - 1.93 A₆₄₇
   Chlorophyll b = 20.36 A₆₄₇ - 5.50 A₆₆₄
   NOTE: Results are expressed in µg mL^-1 of extract, which are multiplied by the extract volume and divided by the number of mL of sample to obtain chlorophyll concentrations in µg mL^-1 of culture.
10. To obtain values in µg chl cell^-1, use the following formula:
    μg chl cell^-1 = µg chl in 1 mL of culture / [cells mL^-1] cells concentration in the culture.

Results

To observe the efficiency of the technique detecting variations in chlorophyll cellular concentration and evaluate the effect of growth factors in C. sorokiniana, a Taguchi L₄ experimental design was established, evaluating CO₂ volume addition, light color, and light intensity. Each factor was assessed at low and high levels, as shown in Table 1, under the conditions defined by the experimental design in Table 2.

Once the experim...

Discussion

The comparative study between H. pluvialis and C. sorokiniana revealed significant differences in chlorophyll production dynamics. While H. pluvialis exhibited a decrease in chlorophyll concentration throughout the experiment, C. sorokiniana showed a steady increase. Additionally, there was initially a lower proportion of chlorophyll a in both species, but this ratio reversed in particular growth conditions, which may give indications of an induction of the production of said ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
C3H6O	Meyer	67-64-1	Acetone 90%
15 mL tube	Biologix	10-9502	Test tube
2510-DTH	Branson	D-73595	Sonicator
5 mL screw cap test tube	Kimax	45066-13100	Test tube
50 mL centrifuge tube	Biologix	10-9151	Test tube
Aluminum foil	Reynolds	611 standard, 12" x 1000 feet	Test tube cover
CaCl2	Meyer	0925-250	Calcium Chloride
Centrifuge	Dynamica	14 R	Centrifuge Refrigerated
CoCl2	Merck	1057-100	Cobalt dichloride
FeCl3	Merck	157740	Iron(III) Chloride
K2HPO4	Meyer	2051-250	Dipotassium Phosphate
KH2PO4	Meyer	2055-250	Monopotassium Phosphate
MgSO4	Meyer	1605-250	Magnesium Sulphate
Micropipette	LabNet	Model Beta-Pette	Micropipette
MnCl2	Merck	429449	Manganese(II) Chloride
Na2 EDTA	Merck	200-449-4	Edatamil, Edetato Disodium Salt Dihydrate
Na2MoO4	Merck	243655	Sodium Molybdate
NaCl	Meyer	2365-500	Sodium Chloride
NaNO3	Meyer	2465-250	Sodium Nitrate
RGB LED stripe	Steren	GAD-LED2	Light source
Spectrophotometer	PerkinElmer	Model Lambda35	Spectrophotometer
spectroradiometer	Gigahertz-Optik	model BTS256
Vortex	Scientific Industries	Vortex-Genie® 2	Vortex
ZnCl2	Merck	208086	Zinc Chloride