Microalgae vs. Higher Plants: A Comparative Analysis of Air Revitalization Efficiency for Advanced Research

Abstract

This article provides a comprehensive scientific analysis for researchers and scientists comparing the air revitalization capabilities of microalgae and higher plants. It explores the foundational biological mechanisms, including photosynthetic efficiency and pollutant metabolism, and examines methodological applications in controlled environments like photobioreactors. The review details common challenges in scaling and optimization, supported by a comparative validation of performance metrics for CO2 fixation, VOC removal, and oxygen production. Synthesizing current research, the article concludes with future directions for strain engineering and clinical implications for improving indoor air quality in biomedical settings.

The Biological Machinery of Air Purification: Unpacking Photosynthetic and Metabolic Pathways

Fundamental Principles of Photosynthesis in Microalgae vs. Higher Plants

Photosynthesis is the fundamental biological process that drives global carbon and oxygen cycles, serving as the primary engine for life on Earth. In the context of developing advanced air revitalization systems for closed environments, understanding the distinct photosynthetic mechanisms of microalgae and higher plants becomes crucial for research and application. While both utilize the core principles of oxygenic photosynthesis—converting light energy, carbon dioxide, and water into chemical energy and oxygen—significant differences exist in their efficiency, adaptability, and implementation potential. Microalgae, unicellular photosynthetic microorganisms, offer remarkable adaptability and efficiency due to their simple structure and aquatic nature [1]. Higher plants, with their complex multicellular organization and specialized tissues, represent the traditional model for terrestrial life support but face different constraints [2]. This guide objectively compares the fundamental principles of photosynthesis in these two distinct biological systems, providing researchers and scientists with experimental data and methodologies relevant for evaluating their application in air revitalization efficiency research.

Core Biological and Structural Differences

The photosynthetic capabilities of microalgae and higher plants are fundamentally shaped by their distinct biological organization and structural adaptations. Microalgae are unicellular, eukaryotic, phototrophic, and heterotrophic microorganisms that thrive in diverse aquatic habitats [1]. Their simple structure lacks true roots, stems, and leaves, with the entire cell body participating directly in photosynthesis. This minimalistic organization allows for highly efficient light and resource utilization. In contrast, higher plants possess complex multicellular structures with specialized photosynthetic (leaves) and non-photosynthetic (roots, stems) tissues. This specialization creates metabolic costs for maintaining non-photosynthetic structures and establishes vascular systems for resource transport between distant organs [3].

The chloroplast architecture also differs significantly between these organisms. In higher plants, thylakoids form elaborate vertical stacks (grana) with helical arrangements, while microalgae typically exhibit simpler thylakoid organization with isolated units or small stacks of up to 10 units [4]. This structural variation influences light harvesting efficiency and photosynthetic regulation.

Table 1: Fundamental Structural and Functional Differences in Photosynthetic Apparatus

Characteristic	Microalgae	Higher Plants
Cellular Organization	Unicellular	Multicellular with tissue specialization
Photosynthetic Structures	Entire cell surface	Specialized leaves with stomata
Structural Support	No roots, stems, or leaves [1]	Complex root and stem systems
Chloroplast Structure	Simpler thylakoid organization [4]	Complex grana stacks with helical thylakoids
Habitat Adaptation	Aquatic environments	Terrestrial environments
Non-Photosynthetic Biomass	Minimal	Significant investment in non-photosynthetic tissues [3]

Comparative Analysis of Photosynthetic Performance

Quantitative assessment of photosynthetic performance reveals distinct advantages and limitations of microalgae versus higher plants, particularly relevant for air revitalization applications where efficiency metrics are critical.

Photosynthetic Efficiency and Growth Rates

Microalgae exhibit superior photosynthetic efficiency, typically reaching over 8% light energy conversion, compared to 1-2% for most traditional crops like sugar cane [1]. This exceptional efficiency translates to significantly faster growth rates, with microalgae growing 20-30% faster than traditional food crops [1]. Their capacity to accumulate large portions of lipids, carbohydrates, proteins, enzymes, and vitamins further enhances their value for integrated life support systems [1].

The light utilization capabilities differ substantially between these organisms. Microalgae possess disproportionately high levels of pigments (chlorophyll, carotenoids, phytochrome proteins) that form extensive "pigment beds," enabling efficient photon capture across broader light spectra [1]. Higher plants primarily utilize chlorophyll a and b within structured antenna complexes, with limited far-red light utilization in most species [5].

Environmental Adaptability and Resource Utilization

Microalgae demonstrate remarkable environmental flexibility, growing efficiently in diverse climate conditions without competing for arable land [1]. Their capacity for both phototrophic and heterotrophic metabolism provides metabolic versatility absent in most higher plants [1]. Experimental studies with Chlorella vulgaris have demonstrated sustained photosynthetic activity and oxygen production even under dynamically cycled temperature conditions (9-27°C), indicating robustness for environmental control systems [6].

Higher plants exhibit more constrained environmental tolerances, with photosynthetic performance highly dependent on maintaining optimal temperature, humidity, and soil conditions. Their gas exchange is governed by stomatal regulation that creates an inherent trade-off between carbon gain and water loss [2]. This hydraulic limitation necessitates complex regulation mechanisms to balance CO₂ uptake against transpiration water loss, especially under water-stressed conditions [2].

Table 2: Quantitative Comparison of Photosynthetic Performance Metrics

Performance Metric	Microalgae	Higher Plants	Experimental Reference
Photosynthetic Efficiency	>8% [1]	1-2% (typical crops) [1]	Laboratory growth analysis
Growth Rate	20-30% faster than traditional crops [1]	Baseline comparison	Biomass accumulation studies
Theoretical Maximum Biomass Yield	~80 g/m²/day or 280 ton/ha/year [3]	Significantly lower	Photobioreactor vs. field studies
CO₂ Sequestration Capacity	1.3 kg CO₂ per kg biomass [7]	Varies by species	Carbon fixation measurements
Temperature Tolerance Range	Broad (e.g., Chlorella: 9-27°C cyclic [6])	Species-dependent, often narrower	Controlled environment experiments
Oxygen Production Rate	C. vulgaris: 0.013-3.15 mgO₂·L⁻¹·h⁻¹ [6]	Varies by species and conditions	Oxygen electrode measurements

Experimental Protocols and Methodologies

Oxygen Production Measurement Under Dynamic Temperature Conditions

Objective: Quantify oxygen production rates of microalgae under temperature cycles simulating spacecraft thermal control systems [6].

Materials:

• Strains: Temperate Chlorella vulgaris and eurythermic Antarctic Chlorophyta
• Photobioreactor System: Temperature-controlled vessel with gas exchange measurement
• Monitoring Equipment: Dissolved oxygen probe, temperature sensor, chlorophyll fluorometer
• Culture Medium: Standard freshwater nutrient medium for Chlorella, modified marine medium for Antarctic strains

Methodology:

• Inoculate microalgae cultures at standardized cell density (OD₆₈₀ ≈ 0.2)
• Apply temperature cycles:
  • C. vulgaris: 9-27°C cycles (28-minute duration)
  • Antarctic Chlorophyta: 4-14°C cycles (28-minute duration)
  • Include constant temperature controls (10°C) for comparison
• Maintain constant light intensity (150-200 μmol photons·m⁻²·s⁻¹) throughout experiment
• Monitor dissolved oxygen concentration at 5-minute intervals using precision oxygen electrode
• Measure chlorophyll fluorescence parameters (Fᵥ/Fₘ) to assess photosynthetic health
• Sample for biomass determination (dry weight) at experiment initiation and conclusion
• Calculate oxygen production rates normalized by volume and biomass

Data Analysis:

• Determine maximum oxygen production rates under cycled vs. constant temperature
• Calculate temperature acclimation index as final O₂ production/initial O₂ production
• Assess statistical significance between conditions using ANOVA with post-hoc testing

Photosynthetic Performance Under Modified Spectral Quality

Objective: Evaluate photosynthetic growth and efficiency of microalgae and plants under simulated M-dwarf starlight spectrum enriched in far-red wavelengths [5].

Materials:

• Test Organisms: Microalgae (Chlorella vulgaris, Dixoniella giordanoi, Microchloropsis gaditana) and plants (Physcomitrium patens, Arabidopsis thaliana)
• Lighting System: Custom lamp system generating M-dwarf spectrum (M7, 365-850 nm), solar spectrum, and far-red monochromatic light (730 nm)
• Growth Chambers: Temperature and humidity-controlled with precise atmospheric regulation
• Analysis Equipment: Spectroradiometer, pulse-amplitude modulation (PAM) fluorometer, biomass quantification tools

Methodology:

• Pre-culture all organisms under standard growth conditions to exponential growth phase
• Transfer to experimental chambers with three light conditions:
  • M7 simulated M-dwarf spectrum
  • SOL simulated solar spectrum
  • FR far-red monochromatic light (730 nm)
• Maintain identical temperature, CO₂ concentration, and nutrient conditions across light treatments
• Measure growth rates via optical density (microalgae) or leaf area/weight (plants) daily
• Assess photosynthetic efficiency using chlorophyll fluorescence parameters (Fᵥ/Fₘ, ΦPSII)
• Conduct light response curves at experiment conclusion to determine photosynthetic capacity
• Analyze pigment composition via HPLC for accessory pigment expression

Data Analysis:

• Calculate specific growth rates under each spectral quality
• Compare photosynthetic efficiency parameters across treatments
• Assess Emerson enhancement effect (synergistic visible + far-red light) by comparing M7 vs. SOL growth

Signaling Pathways and Photosynthetic Mechanisms

The core photosynthetic pathways share fundamental similarities between microalgae and higher plants, but significant differences exist in their regulatory mechanisms and light-harvesting strategies.

Electron Transport and Carbon Fixation

Both microalgae and higher plants utilize linear electron transport (LET) involving photosystem II (PSII) and photosystem I (PSI) in sequence. The process begins with photon capture by light-harvesting complexes, excitation energy transfer to reaction centers (P680 in PSII, P700 in PSI), and photochemical charge separation [3]. Water splitting at the manganese-containing complex in PSII generates protons, electrons, and molecular oxygen [8].

Electrons travel through the electron transport chain via plastoquinone (PQ), cytochrome b₆f complex, and plastocyanin to PSI, where further excitation drives NADP⁺ reduction to NADPH via ferredoxin [3]. The proton gradient established across the thylakoid membrane drives ATP synthesis through ATP synthase [8].

Microalgae exhibit more flexible electron transport pathways, with significant cyclic electron flow (CEF) around PSI mediated by PGR5 and PGRL1 proteins, enhancing photoprotection and optimizing ATP/NADPH stoichiometry for metabolism [3].

Photoprotective Mechanisms and Light Acclimation

Microalgae possess dynamic photoprotective strategies, including non-photochemical quenching (NPQ) mediated by light-harvesting complex stress-related (LHCSR) proteins [3]. When exposed to excessive light, microalgae rapidly induce energy dissipation mechanisms through xanthophyll cycle pigments (violaxanthin, zeaxanthin) [3].

Higher plants utilize similar xanthophyll cycle-based photoprotection but rely primarily on PSII protein phosphorylation and state transitions for excess energy management. The hydraulic constraints of terrestrial environments have led to sophisticated stomatal regulation mechanisms that balance CO₂ uptake against water loss, creating an intrinsic trade-off between carbon gain and hydraulic risk [2].

Research Reagent Solutions and Essential Materials

For researchers investigating photosynthetic performance in microalgae and higher plants, particularly for air revitalization applications, the following reagents and materials are essential:

Table 3: Essential Research Reagents and Experimental Materials

Reagent/Material	Function/Application	Example Use Cases
PAM Fluorometer	Measures chlorophyll fluorescence parameters (Fᵥ/Fₘ, ΦPSII)	Quantifying photosynthetic efficiency and photoinhibition [5]
Dissolved Oxygen Probe	Precise measurement of oxygen evolution rates	Quantifying air revitalization capacity [6]
Custom Spectral Lighting	Simulates specific light environments (M-dwarf, far-red enriched)	Testing photosynthesis under non-terrestrial spectra [5]
Temperature-Controlled Photobioreactors	Maintains precise temperature regimes during growth studies	Investigating thermal tolerance and performance [6]
Chlorella vulgaris Strains	Model microalga for photosynthetic research	Baseline oxygen production studies [6]
Antarctic Chlorophyta Strains	Eurythermic microalgae for extreme environment adaptation	Low-temperature photosynthesis studies [6]
Arabidopsis thaliana	Model plant for comparative photosynthesis studies	Reference for terrestrial plant responses [5]
Nutrient Media Formulations	Standardized growth media for consistent cultivation	Maintaining optimal growth conditions across experiments
HPLC Pigment Analysis System	Separation and quantification of photosynthetic pigments	Assessing photosynthetic apparatus composition and acclimation [5]

Microalgae and higher plants represent two evolutionarily distinct approaches to oxygenic photosynthesis with compelling contrasts in efficiency, adaptability, and implementation potential for air revitalization systems. Microalgae demonstrate superior photosynthetic efficiency (over 8%), faster growth rates, and remarkable environmental flexibility, making them exceptionally promising for compact, efficient air revitalization in controlled environments [1]. Their simple cellular structure, capacity for both phototrophic and heterotrophic growth, and tolerance to dynamic temperature conditions provide significant advantages for engineered life support systems [6]. Higher plants offer complementary benefits through their more complex ecosystem integration, production of diverse secondary metabolites, and psychological value in human habitats, albeit with lower photosynthetic efficiency and greater resource requirements [2].

The experimental data and methodologies presented provide researchers with standardized approaches for quantitative comparison of these biological systems. Future research directions should focus on harnessing the superior efficiency of microalgae while integrating the complementary benefits of plant-based systems, potentially through hybrid approaches that optimize the unique strengths of each photosynthetic strategy for advanced air revitalization applications.

Comparative Analysis of Carbon Concentration Mechanisms (CCMs) and Rubisco Efficiency

Carbon Concentration Mechanisms (CCMs) represent a critical evolutionary adaptation in photosynthetic organisms, enhancing the efficiency of the central carbon-fixing enzyme, Ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco). This comparative guide examines the fundamental differences in CCM strategies between microalgae and higher plants, with specific application to air revitalization efficiency research. Rubisco's catalytic inefficiency, characterized by a slow turnover rate and competitive oxygenation reaction that leads to photorespiration, has driven the evolution of diverse biological solutions to concentrate CO₂ around the enzyme's active site [9] [10]. Understanding these mechanisms is paramount for optimizing photosynthetic efficiency in both agricultural and controlled environmental systems, including bioregenerative life support systems for space applications [11].

Microalgae, comprising both prokaryotic cyanobacteria and eukaryotic green algae, have evolved sophisticated biophysical CCMs involving subcellular compartmentalization. These organisms actively transport inorganic carbon (Ci) and compartmentalize Rubisco within specialized structures like carboxysomes and pyrenoids, creating CO₂-rich microenvironments that suppress photorespiration [7] [12]. In contrast, higher plants have primarily evolved biochemical CCMs, such as the C₄ and Crassulacean Acid Metabolism (CAM) pathways, which spatially or temporally separate carbon fixation from the Calvin cycle [13]. C₃ plants, which lack specialized CCMs, rely solely on the diffusion of atmospheric CO₂ and represent the ancestral photosynthetic pathway [14].

This analysis provides researchers with a structured comparison of CCM operational principles, quantitative performance metrics, and experimental approaches for evaluating carbon fixation efficiency. The findings have significant implications for engineering enhanced carbon capture in crops and developing efficient photobioreactor systems for simultaneous air revitalization and biomass production in closed environments [15] [11].

Fundamental Mechanisms of Carbon Concentration

Microalgal Biophysical CCMs

Microalgae employ sophisticated biophysical CCMs that operate through three coordinated mechanisms: active inorganic carbon transport across membranes, carbonic anhydrase-mediated conversion between CO₂ and bicarbonate (HCO₃⁻), and spatial compartmentalization of Rubisco [7]. The foundational structure of this system in green algae like Chlamydomonas reinhardtii involves a pyrenoid—a specialized, starch-coated microcompartment within the chloroplast where Rubisco is densely packed [14].

The CCM operates through two potential modes validated by chloroplast-level modeling [14]. In the active uptake mode, energy-dependent HCO₃⁻ pumps transport bicarbonate across the chloroplast envelope against a concentration gradient (Fig. 1). The accumulated stromal HCO₃⁻ diffuses into the thylakoid lumen, where it traverses the pyrenoid. Within the pyrenoid matrix, carbonic anhydrase (CA) rapidly converts HCO₃⁻ to CO₂, creating a localized high-CO₂ environment that saturates Rubisco and minimizes oxygenation. A critical component is the diffusion barrier, comprising stacked thylakoid membranes and a starch sheath, which reduces CO₂ leakage and prevents futile cycling [14].

In the passive uptake mode, CO₂ diffuses freely across the chloroplast envelope into the stroma, where stromal CA hydrates it to HCO₃⁻. The alkaline pH of the stroma favors HCO₃⁻ accumulation, which then follows the same pathway into the pyrenoid for concentration and fixation. This mode operates with remarkable energetic efficiency of 2-3 ATPs per CO₂ fixed when combined with effective diffusion barriers [14].

Cyanobacteria employ a similar strategy using carboxysomes—icosahedral protein microcompartments that co-encapsulate Rubisco and CA. Bicarbonate is actively transported into the cell and diffuses into carboxysomes, where CA converts it to CO₂, creating a concentrated carbon environment around Rubisco [12].

Figure 1: Comparative schematic of Carbon Concentration Mechanisms in microalgae and C₄ plants. Microalgae utilize subcellular compartmentalization in pyrenoids, while C₄ plants employ spatial separation between mesophyll and bundle sheath cells.

Plant Biochemical CCMs

In contrast to microalgal biophysical approaches, C₄ plants like maize and sorghum evolved a biochemical CCM that spatially separates initial carbon fixation from the Calvin cycle [13]. This mechanism involves two distinct photosynthetic cell types: mesophyll cells and bundle sheath cells arranged in concentric layers around vascular tissue (Kranz anatomy).

In mesophyll cells, the enzyme phosphoenolpyruvate (PEP) carboxylase initially fixes HCO₃⁻ into four-carbon organic acids (malate or aspartate). These C₄ acids then diffuse to bundle sheath cells where they are decarboxylated, releasing CO₂ in close proximity to Rubisco. The concentrated CO₂ suppresses photorespiration, while the three-carbon residue returns to the mesophyll cells to regenerate PEP, completing the cycle [13].

The crassulacean acid metabolism (CAM) pathway represents a temporal rather than spatial separation of carbon fixation. CAM plants, typically adapted to arid environments, open their stomata at night to fix CO₂ into organic acids, which are stored in vacuoles. During the day, when stomata are closed to reduce water loss, these acids are decarboxylated, providing CO₂ for Rubisco and the Calvin cycle [13].

C₃ plants like wheat, rice, and soybeans lack specialized CCMs and rely solely on atmospheric CO₂ diffusion. Their Rubisco therefore operates in a photorespiratory environment where the oxygenase reaction competes with carboxylation, especially under conditions of high temperature, light intensity, or water stress [12] [14].

Quantitative Performance Comparison

Table 1: Comparative Efficiency Metrics of CCM Systems in Photosynthetic Organisms

Parameter	C₃ Plants	C₄ Plants	CAM Plants	Microalgae	Measurement Context
Rubisco Carboxylation Rate (kcat_c, s⁻¹)	2-5 [12]	3-6 [13]	2-4 [13]	5-15 [9] [10]	Per active site at 25°C
CO₂ Concentration at Rubisco Site (μM)	5-15 [14]	50-200 [13]	30-100 [13]	100-500 [14]	Estimated from modeling
Photorespiration Rate Relative to Carboxylation	20-40% [12]	3-5% [13]	5-15% [13]	2-10% [7]	At 25°C, ambient CO₂
Energy Cost (ATP/CO₂ fixed)	3 [14]	5 [13]	4.5-6.5 [13]	2-4 [14]	Includes CCM operation
Carbon Sequestration Efficiency	Baseline	10-50% higher than C₃ [13]	Variable	10-50× terrestrial plants [7]	Per unit biomass
Rubisco Specificity Factor (SC/O)	80-100 [13]	70-85 [13]	75-90 [13]	50-200 (varies by species) [10]	Relative specificity for CO₂ vs O₂

Table 2: Air Revitalization Performance in Controlled Systems

Parameter	C₃ Plants	C₄ Plants	Microalgae	Notes
CO₂ Removal Rate (mg CO₂/g biomass/h)	1.5-3.5 [11]	2.5-5.0 [11]	8-15 [11]	At 2000 ppm CO₂, continuous light
O₂ Production (mg O₂/g biomass/h)	1.1-2.5 [11]	1.8-3.6 [11]	6-11 [11]	Coupled to CO₂ fixation
Water Consumption (L/kg biomass)	500-1000 [11]	250-500 [11]	50-200 (recycled) [11]	Includes transpiration/evaporation
Biomass Productivity (g/m²/day)	10-25 [11]	20-40 [11]	80-200 [11]	Optimized cultivation conditions
System Footprint (m²/person)	20-40 [11]	15-25 [11]	5-10 [11]	For complete air revitalization

The quantitative comparison reveals distinct advantages of microalgal systems for air revitalization applications. Microalgae exhibit superior CO₂ fixation and O₂ production rates per unit biomass, significantly reduced water requirements through recycling, and substantially higher biomass productivity, resulting in a smaller system footprint for supporting human life [11]. These advantages stem from their efficient biophysical CCMs, which concentrate CO₂ around Rubisco with lower energy costs compared to biochemical CCMs in C₄ plants [14].

The evolutionary trade-offs in Rubisco kinetics are particularly revealing. C₃ plants exhibit higher Rubisco specificity factors (SC/O), reflecting adaptation to lower CO₂ environments, while C₄ plants and microalgae operating with CCMs can utilize Rubisco variants with higher catalytic rates but lower specificity [13]. Microalgal Rubisco demonstrates the widest variation in specificity factors, indicating diverse evolutionary adaptations to different ecological niches and CCM configurations [10].

Rubisco Engineering and Synthetic CCM Approaches

Directed Evolution of Rubisco

Recent advances in protein engineering have demonstrated the potential to enhance Rubisco's catalytic properties through directed evolution. MIT chemists successfully improved the efficiency of a bacterial Rubisco using a continuous evolution platform called MutaT7, which allows for rapid mutagenesis and screening in living cells [9]. Through six rounds of directed evolution, they identified three mutations near the enzyme's active site that improved oxygen resistance and increased catalytic efficiency by up to 25% [9].

This approach represents a significant advancement over traditional error-prone PCR methods, which typically introduce only one or two mutations per generation and generate smaller mutant libraries. The MutaT7 system enables higher mutation rates and continuous evolution under selective pressure, allowing researchers to explore a broader sequence space for beneficial mutations [9] [10]. The improved Rubisco variants showed reduced oxygenation activity, preferentially reacting with carbon dioxide even in oxygen-rich environments [9].

Synthetic Carbon Concentrating Mechanisms

Engineering synthetic CCMs into C₃ crops represents a promising approach to enhance photosynthetic efficiency. Recent breakthrough research has demonstrated the reprogramming of bacterial encapsulins into modular carbon-fixing nanocompartments [12]. These synthetic microcompartments from Quasibacillus thermotolerans (QtEnc) can be loaded with diverse Rubisco isoforms by fusing a short cargo-loading peptide (CLP) to the enzyme [12].

The structural configuration of these synthetic nanocompartments enables targeted encapsulation of multiple Rubisco forms while preserving catalytic activity. For Form I Rubiscos from tobacco (Nicotiana tabacum) and the bacterium Rhodobacter sphaeroides, researchers appended the CLP tag to the C-terminus of the RbcS small subunit, which minimized disruption to catalytic function [12]. This modular design establishes a foundation for creating plant-compatible synthetic carboxysomes that could potentially enhance CO₂ concentration around Rubisco in C₃ plants [12].

Figure 2: Engineering strategies for enhancing carbon fixation. Two primary approaches include directed evolution of Rubisco kinetics and creation of synthetic carbon concentrating compartments through encapsulin reprogramming.

Experimental Methods for CCM Analysis

Rubisco Activity Assays

Quantifying Rubisco catalytic parameters requires specialized biochemical assays to determine carboxylation velocity, substrate affinity, and specificity. The radiometric ^14CO₂ fixation assay remains the gold standard for measuring Rubisco carboxylation rates [10]. This method involves incubating purified Rubisco with radiolabeled ^14CO₂ and ribulose-1,5-bisphosphate (RuBP) for precise time intervals before terminating the reaction with acid. The acid-stable ^14C-labeled products are then quantified by scintillation counting [10].

For high-throughput screening of Rubisco variants, a novel 3-phosphoglycerate (3PG) biosensing approach has been developed [10]. This system links Rubisco activity to transcription of a reporter protein and quantifies intracellular Rubisco concentration, enabling normalization of carboxylation activity by enzyme abundance. This method facilitates rapid identification of high-performing Rubisco homologs from natural sources and directed evolution campaigns [10].

The specificity factor (SC/O), which quantifies Rubisco's ability to discriminate between CO₂ and O₂, is determined by simultaneously measuring carboxylase and oxygenase activities using either mass spectrometry or an oxygen electrode system [13]. This parameter is typically assessed under controlled atmospheric conditions with precise O₂:CO₂ ratios.

In Vivo Carbon Fixation Analysis

Rubisco-dependent E. coli (RDE) strains provide a powerful selection system for evaluating Rubisco function in vivo [10]. These engineered bacteria lack native phosphoribulokinase (Prk) and become dependent on heterologously expressed Rubisco for growth by converting ribulose bisphosphate (RuBP) into metabolic intermediates. Recent improvements to this system include deletion of ribose 5-phosphate isomerase (rpi) to force pentose phosphate pathway flux through RuBP, creating stronger coupling between Rubisco activity and cellular fitness [10].

For evaluating complete CCM function, membrane inlet mass spectrometry (MIMS) enables precise measurement of CO₂ and O₂ fluxes in intact cells or isolated organelles. This technique allows researchers to quantify inorganic carbon uptake, photosynthetic oxygen evolution, and photorespiratory activity simultaneously [14]. When combined with isotopic labeling with ^13C or ^18O, MIMS can trace carbon flow through different metabolic pathways and evaluate CCM efficiency under various environmental conditions.

Chlorophyll fluorescence imaging provides a non-invasive method to monitor photosynthetic efficiency and photorespiration in vivo. Parameters such as quantum yield of photosystem II (ΦPSII) and non-photochemical quenching (NPQ) can indicate CCM functionality, particularly under carbon-limiting conditions [14].

Research Reagents and Tools

Table 3: Essential Research Reagents for CCM and Rubisco Studies

Reagent/Tool	Application	Key Features	Experimental Considerations
Rubisco-Dependent E. coli (RDE) Strains [10]	In vivo selection of functional Rubisco variants	Couples Rubisco activity to bacterial growth	Requires careful control of expression levels; potential for false positives from solubility mutations
MutaT7 Continuous Evolution System [9]	Directed evolution of Rubisco	Enables rapid mutagenesis and screening in living cells	Allows exploration of larger mutational space than error-prone PCR
3PG Biosensor System [10]	High-throughput screening of Rubisco activity	Links carboxylation to reporter gene expression; normalizes by enzyme concentration	Enables quantitative screening without radioactive materials
^14C-Labeled Sodium Bicarbonate [10]	Radiometric Rubisco activity assays	Gold standard for carboxylation rate measurement	Requires radiation safety protocols; specialized detection equipment
Encapsulin Nanocompartments (QtEnc) [12]	Synthetic CCM engineering	Self-assembling protein compartments; modular cargo loading	CLP tagging position critical for preserving Rubisco activity
Anti-Rubisco Antibodies [12]	Rubisco quantification and localization	Species-specific antibodies available	Important for normalizing activity measurements to enzyme concentration
Membrane Inlet Mass Spectrometry (MIMS) [14]	Gas exchange measurements in intact cells	Simultaneous monitoring of O₂ and CO₂ fluxes	Enables real-time analysis of CCM function under varying conditions

This comparative analysis reveals fundamental differences in carbon concentration strategies between microalgae and higher plants, with significant implications for air revitalization applications. Microalgae employ biophysical CCMs based on subcellular compartmentalization in pyrenoids, creating high-CO₂ microenvironments around Rubisco through active transport and diffusion barriers. In contrast, C₄ plants utilize biochemical CCMs that spatially separate carbon fixation from the Calvin cycle, while C₃ plants lack specialized concentrating mechanisms entirely [7] [14] [13].

The quantitative performance data demonstrates clear advantages of microalgal systems for bioregenerative life support, with superior CO₂ fixation rates (8-15 mg CO₂/g biomass/h), oxygen production capacity (6-11 mg O₂/g biomass/h), and significantly reduced water requirements compared to terrestrial plants [11]. These characteristics, combined with their rapid growth rates and ability to thrive in closed systems, position microalgae as optimal candidates for air revitalization in controlled environments.

Recent breakthroughs in Rubisco engineering through directed evolution [9] and synthetic CCM development using encapsulin nanocompartments [12] provide promising pathways for enhancing carbon fixation efficiency in both agricultural and specialized applications. The experimental frameworks and research tools outlined in this analysis will support continued innovation in this critical field, potentially enabling the development of next-generation biological systems for carbon capture and atmospheric regeneration.

The quest for efficient biological air revitalization systems has intensified with growing concerns over environmental pollution and human health. Within this context, microalgae and higher plants represent two fundamental biological systems capable of metabolizing airborne pollutants, including volatile organic compounds (VOCs) and particulate matter. While higher plants have been traditionally studied for phytoremediation applications, microalgae—photosynthetic microorganisms inhabiting aquatic and moist terrestrial environments—demonstrate remarkable metabolic versatility and degradation efficiency that warrants detailed scientific comparison [16] [17]. This review systematically compares the metabolic pathways, degradation efficiencies, and experimental protocols for pollutant remediation by these two biological systems, providing researchers with quantitative data to inform biotechnological development and environmental application.

Microalgae possess several distinctive advantages for pollutant degradation, including rapid growth rates, high surface-area-to-volume ratios, and diverse metabolic capabilities that can be optimized through cultivation conditions [18] [15]. Their emission of VOCs itself constitutes a sophisticated biological response mechanism to environmental stressors, functioning as infochemicals in aquatic ecosystems and offering protective roles against abiotic stresses [17]. This review objectively analyzes the current scientific understanding of these systems, with particular emphasis on comparative performance metrics and methodological approaches for evaluating degradation efficiency.

Metabolic Pathways for VOC and Particulate Matter Degradation

Microalgae Metabolic Pathways

Microalgae employ multiple interconnected metabolic pathways for pollutant transformation and degradation, with significant variations between species and environmental conditions. The primary pathways involve direct enzymatic transformation, biosorption, bioaccumulation, and biodegradation processes [18] [19].

Terpenoid Synthesis and Carotenoid Degradation: Microalgae synthesize terpenoids via two principal pathways: the methylerythritol-4-phosphate (MEP) pathway in plastids for isoprene and monoterpene production, and the mevalonate (MVA) pathway for sesquiterpenes [17]. The MEP pathway utilizes pyruvate and glyceraldehyde-3-phosphate as initial substrates, proceeding through multiple enzymatic steps to dimethylallyl pyrophosphate (DMAPP)—the immediate precursor for isoprene and monoterpenes. Carotenoid degradation represents another significant source of VOC production in microalgae, generating compounds such as β-cyclocitral, β-ionone, and geranylacetone through oxidative cleavage reactions [17]. These compounds function as allelopathic agents and stress response molecules in aquatic environments.

Fatty Acid Oxidation and Halogenated Compound Formation: The oxidative degradation of fatty acids leads to the production of C6 green leaf volatiles (GLVs), including alcohols and aldehydes, which increase under stress conditions such as high temperature [17]. Additionally, many marine microalgae produce halogenated hydrocarbons through haloperoxidase enzymes that catalyze hydrogen peroxide-mediated oxidation of halide ions. This process is light-dependent, with elevated production rates observed under high light intensity due to increased reactive oxygen species (ROS) generation [17].

Nutrient Stress Response Pathways: Under phosphorus or nitrogen limitation, microalgae significantly upregulate VOC emission through multiple metabolic adjustments. Non-nitrogen conditions induce overexpression of genes encoding pyruvate kinase, malic enzyme, phosphotransacetylase, and aspartate aminotransferase—key enzymes involved in producing precursors for terpenoid and benzenoid synthesis [17]. This transcriptional regulation enhances flux through both the MEP and shikimate pathways, substantially increasing VOC diversity and emission rates under nutrient stress.

The following diagram illustrates the interconnected metabolic pathways for VOC production in microalgae under various environmental conditions:

Higher Plant Metabolic Pathways

Higher plants employ fundamentally different structural and metabolic strategies for air revitalization, primarily utilizing leaf surface structures and internal metabolic pathways. The primary mechanisms include:

Stomatal Uptake and Internal Transformation: Gaseous pollutants enter plant tissues primarily through stomata, followed by dissolution in apoplastic water and diffusion into cells. Particulate matter is primarily intercepted on leaf surfaces based on morphology (hairs, ridges, waxes) with limited internalization. Once internalized, organic pollutants undergo enzymatic transformation through cytochrome P450 monooxygenases, peroxidases, and transferases, leading to conjugation with glutathione, sugars, or amino acids, followed by compartmentalization in vacuoles or cell walls.

Rhizosphere Interactions: Higher plants additionally leverage root-associated microbial communities for extended degradation capabilities, particularly in the soil ecosystem. This plant-microbe partnership significantly expands the metabolic range for pollutant degradation beyond the plant's native enzymatic capabilities.

Comparative Limitations: Unlike microalgae, higher plants generally exhibit slower metabolic response times to environmental pollutants and more limited capabilities for degrading complex organic contaminants due to their more specialized metabolic networks.

Quantitative Comparison of Pollutant Degradation Efficiency

Direct comparative studies between microalgae and higher plants for air revitalization are limited in the current literature, as most research focuses on their respective applications in different environments (aquatic vs. terrestrial). However, extrapolation from wastewater treatment studies and metabolic efficiency analyses provides valuable insights into their relative capabilities for pollutant degradation.

Table 1: Comparative Pollutant Removal Efficiencies of Microalgae and Higher Plants

Pollutant Category	Specific Pollutant	Microalgae Efficiency	Higher Plants Efficiency	Notes
Nutrients	Total Nitrogen (TN)	21.3–44.3% [20]	Below 20% [20]	Microalgae show superior direct nutrient removal
	Total Phosphorus (TP)	53.3–80.0% [20]	~10% [20]	Microalgae outperform plants in phosphorus assimilation
Organic Matter	Chemical Oxygen Demand (COD)	Up to 98.8% [20]	Variable (species-dependent)	Attached microalgae systems show exceptional performance
Emerging Contaminants	Antibiotics (e.g., SMZ)	Significant removal via biodegradation [20]	Limited data	Microalgae show specialized degradation pathways
	Heavy Metals	45-65% of BOD/COD [18]	Limited direct comparison	Microalgae employ biosorption, bioaccumulation
Production Benefits	Biomass Yield	0.22–1.81 g L−1 [18]	Lower biomass per area	Microalgae enable valuable byproduct generation
	Oxygen Release	Enhanced through photosynthesis [20]	Standard photosynthetic rates	Microalgae increase DO for improved nitrification

Table 2: Microalgae Performance in Targeted Pollutant Removal

Microalgae Species	Target Pollutant	Removal Efficiency	Experimental Conditions
Chlorella variabilis	Domestic wastewater nutrients	1.72 g L−1 biomass production [18]	Domestic wastewater cultivation
Scenedesmus abundans	Domestic wastewater contaminants	3.55 g L−1 biomass production [18]	Domestic wastewater cultivation
Scenedesmus sp.	Municipal wastewater	1.81 g L−1 biomass production [18]	Municipal wastewater application
Chlorella sorokiniana	Polyethylene microplastics	IC50 of 100 mg/L [21]	96h exposure in BBM medium
C. pyrenoidosa	Heavy metals (Hg, Ag)	>50% removal [22]	Domestic wastewater
C. pyrenoidosa	Pharmaceutical (clarithromycin)	~80% removal [22]	Controlled laboratory conditions
Chlorella vulgaris-Scenedesmus quadricauda-Arthrospira platensis consortium	Pesticide (malathion)	Up to 99% removal [22]	Urban wastewater testing

The quantitative data clearly demonstrates microalgae's superior efficiency in nutrient removal and biomass production compared to higher plants. This advantage stems from their direct assimilation capabilities and diverse enzymatic machinery for pollutant transformation. Additionally, microalgae systems offer the valuable advantage of generating harvestable biomass for biofuel, feed, or biochemical production—creating a circular economy approach to pollution mitigation [15].

Experimental Protocols for Pollutant Degradation Assessment

Microalgae Cultivation and Exposure Systems

Standardized protocols for evaluating pollutant degradation by microalgae require careful control of cultivation parameters and exposure conditions:

Photobioreactor Setup: For VOC degradation studies, closed photobioreactor systems (0.5-5L working volume) with precise environmental control are recommended. Optimal conditions typically include: temperature maintained at 25±2°C [20], continuous illumination at 60-200 μmol photons m⁻² s⁻¹ using cool white fluorescent lamps, and mixing provided by air bubbling (0.22 μm filtered air) at 0.5-1 vvm (volume per volume per minute) [21]. The pH should be maintained at 6.8-7.2 using CO₂ supplementation or buffer systems as needed.

Experimental Design for Toxicity Assessment: For determining half-maximal inhibitory concentrations (IC50), prepare a concentration series of the target pollutant (e.g., 0-150 mg/L for microplastics [21]). Inoculate triplicate vessels with mid-exponential phase microalgae cultures (initial biomass concentration 0.1-0.3 g/L). Monitor growth kinetics for 96 hours for IC50 determination or through full growth cycle (typically 14 days) for comprehensive degradation analysis [21].

Attached Microalgae Systems: For tidal flow constructed wetlands simulating in situ conditions, configure systems with bed filler material (e.g., quartz sand, Φ=4-8 mm) and inoculate with activated sludge to establish diverse microbial communities. Operate with alternating tidal cycles (e.g., 6 hours flooding, 6 hours rest) to optimize oxygen transfer and nutrient distribution [20].

The following workflow diagram outlines a standardized experimental approach for assessing microalgae-based pollutant degradation:

Analytical Methods for Degradation Assessment

Comprehensive evaluation of pollutant degradation requires multiple analytical approaches:

Growth and Biomass Analysis: Monitor algal growth daily using optical density (OD680-750), cell counting with hemocytometer, or chlorophyll fluorescence (Fv/Fm). Harvest biomass at stationary phase for dry weight determination (filtering through pre-weighed glass fiber filters, drying at 105°C to constant weight) [21].

Biochemical Composition Analysis: Quantify pigment content (chlorophyll a, b, carotenoids) by solvent extraction (90% acetone or DMSO) and spectrophotometric measurement using established equations [21]. Analyze lipid content gravimetrically after extraction (Bligh & Dyer method), protein content by Lowry or Bradford assay, and carbohydrate content by phenol-sulfuric acid method [21].

VOC Collection and Analysis: Collect VOCs using sorbent tubes (Tenax TA, Carbograph) with low-flow sampling pumps (10-50 mL/min). Analyze via thermal desorption coupled with gas chromatography-mass spectrometry (TD-GC-MS) with database matching (NIST, Wiley libraries) [16]. For high-resolution analysis, employ comprehensive two-dimensional GC×GC-TOF-MS.

Pollutant-Specific Analysis: For emerging contaminants like antibiotics, utilize liquid chromatography with tandem mass spectrometry (LC-MS/MS) for quantification [20]. For microplastics, employ microscopy (SEM), Fourier-transform infrared spectroscopy (FT-IR), and micro-FT-IR for chemical mapping [21].

Oxidative Stress Markers: Quantify reactive oxygen species (ROS) using fluorescent probes (DCFH-DA), measure antioxidant enzyme activities (SOD, CAT, APX), and analyze non-enzymatic antioxidants (phenolics, flavonoids) to assess cellular stress responses [21].

The Scientist's Toolkit: Essential Research Reagents and Materials

Table 3: Essential Research Reagents for Microalgae Pollutant Degradation Studies

Category/Reagent	Specification	Application/Function	Representative Examples
Microalgae Strains	Axenic cultures, validated identity	Pollutant degradation studies	Chlorella vulgaris, Scenedesmus abundans, Chlorella sorokiniana [18]
Culture Media	Standardized formulations	Optimized growth support	Bold's Basal Medium (BBM), BG-11, WC medium [21]
Pollutant Standards	Analytical grade, certified reference materials	Exposure studies quantification	Sulfamethazine (SMZ), polyethylene microplastics, heavy metal standards [20] [21]
Analytical Sorbents	High purity, thermal stability	VOC collection and pre-concentration	Tenax TA, Carbograph, mixed-bed sorbent tubes [16]
Extraction Solvents	HPLC/GC grade, low background	Metabolite and pollutant extraction	Acetone, methanol, dichloromethane, n-hexane [21]
Biochemical Assay Kits	Validated protocols, standardized	Cellular component quantification	Lipid extraction kits, protein assay kits, carbohydrate assay kits [21]
Molecular Biology Reagents	Molecular grade, high purity	Gene expression analysis	RNA extraction kits, cDNA synthesis kits, qPCR reagents [17]
Microscopy Supplies	Specific membrane filters	Cell observation and enumeration	Glass fiber filters, polycarbonate membrane filters [21]

Microalgae demonstrate clear advantages over higher plants in pollutant degradation efficiency, particularly for nutrients, emerging contaminants, and complex organic pollutants. Their rapid growth, diverse metabolic capabilities, and adaptability to various cultivation systems position them as superior candidates for advanced air and water revitalization applications. The well-characterized metabolic pathways for VOC production and degradation in microalgae provide a robust foundation for biotechnological optimization.

Significant research gaps remain in standardizing performance metrics between aquatic and terrestrial systems, optimizing hybrid microalgae-based treatment technologies [23], and developing commercial-scale applications that leverage microalgae's full potential for simultaneous environmental remediation and biomass valorization. Future research should prioritize integrating multi-omics approaches to elucidate degradation pathways, engineering optimized cultivation systems for enhanced gas exchange, and developing economic models that capitalize on the circular bioeconomy potential of microalgae-based pollution control systems.

This guide provides an objective comparison of the anatomic and structural features of microalgae and higher plants, with a specific focus on biomass distribution and surface area, and their direct impact on performance for air revitalization efficiency. It is structured to support researchers and scientists in the field by presenting consolidated experimental data, detailed methodologies, and essential research tools.

The following table summarizes the core anatomic and structural advantages of microalgae that underpin their superior performance in gas exchange and biomass productivity per unit area compared to higher plants.

Feature	Microalgae	Higher Plants (Typical Terrestrial Crops)	Impact on Air Revitalization Efficiency
Photosynthetic Surface Area	Entire cell surface exposed to medium; up to 100x greater surface area-to-volume ratio [24].	Limited to leaf surface area; significant non-photosynthetic structures (stems, roots) [24].	Microalgae achieve far more efficient contact between photosynthetic apparatus and air/medium.
Biomass Distribution	Unicellular or simple multicellular; virtually all cells contribute to photosynthesis and gas exchange [25].	Complex differentiation into photosynthetic (leaf) and non-photosynthetic (root, stem) tissues [24].	A larger proportion of microalgal biomass is directly dedicated to CO₂ capture and O₂ production.
Architectural Complexity	Simple, non-vascular structure; direct diffusion of gases [25].	Vascular systems required to transport gases and nutrients; introduces inefficiencies [24].	Eliminates internal resistance and energy costs associated with gas transport through complex tissues.
Carbon Sequestration Rate	High: 1.0 - 3.7 g CO₂/L/day reported in optimized photobioreactors [26].	Lower: Terrestrial ecosystems absorb ~30% of anthropogenic CO₂, but efficiency is reducing with climate change [24].	Microalgae systems can be designed for significantly higher volumetric CO₂ fixation rates.
Growth Rate & Biomass Yield	Rapid doubling; high biomass productivity per unit area; lipid content up to 60-70% dry weight in some strains [26] [27] [28].	Slower growth; lower biomass yield per unit area and time [27].	Enables faster biomass generation and valuable compound production in a smaller footprint.

Experimental Protocols for Performance Comparison

To objectively compare the air revitalization potential of microalgae and higher plants, researchers typically quantify key physiological and growth parameters. Below are detailed methodologies for core experiments.

Experiment 1: Quantifying Carbon Sequestration Rate

This protocol measures the direct CO₂ fixation efficiency of a system, a critical metric for air revitalization.

• Objective: To determine the rate of carbon dioxide uptake by microalgae or higher plants in a controlled environment.
• Materials:
  • For Microalgae: Photobioreactor (e.g., tubular, flat-panel), CO₂ cylinder with regulator, pH meter, dissolved CO₂ probe, microalgae culture (e.g., Chlorella sp., Chlorococcum sp.), defined growth medium (e.g., BBM, TAP) [29] [24] [25].
  • For Higher Plants: Sealed plant growth chamber, CO₂ sensor, potted plant specimen (e.g., lettuce, Arabidopsis), soil or growth substrate.
• Methodology:
  • System Setup & Inoculation: Set up the photobioreactor or plant chamber. Inoculate the photobioreactor with a known density of microalgae (e.g., 2 × 10⁷ cells mL⁻¹) [29]. For plants, use a uniformly sized specimen.
  • CO₂ Injection & Monitoring: Introduce a known concentration of CO₂ into the system. For microalgae, this is often a continuous flow (e.g., air enriched with 1-5% CO₂) [25]. Continuously monitor and log the decrease in CO₂ concentration within the headspace or the dissolved CO₂ in the medium over time.
  • Biomass Measurement: At the end of the experiment (e.g., 24-72 hours), harvest the biomass. For microalgae, measure dry weight [30]. For plants, measure fresh and dry weight of the aerial parts.
  • Calculation: The carbon sequestration rate can be calculated from the rate of CO₂ depletion from the atmosphere. Alternatively, for microalgae, the fixed carbon can be estimated from the biomass productivity, knowing that ~1.7-2.0 kg of CO₂ is required to produce 1 kg of algal biomass [25].
• Data Output: Carbon fixation rate expressed as g CO₂/L/day (for microalgae) or g CO₂/m²/day (for plants).

Experiment 2: Analyzing Biomass Productivity and Composition

This protocol assesses the growth rate and the distribution of valuable compounds within the biomass, indicating the efficiency of carbon utilization.

• Objective: To measure the growth rate and biochemical composition (e.g., lipids, proteins) of microalgae and higher plants.
• Materials:
  • Spectrophotometer, centrifuge, oven/freeze dryer, lipid extraction apparatus (e.g., Soxhlet), Bradford assay kit, microalgae/higher plant samples [27] [28] [30].
• Methodology:
  • Growth Monitoring: For microalgae, track growth daily by measuring optical density (OD) at 680 nm and cell count using a hemocytometer or automated cell counter [31] [30]. For plants, measure leaf area, stem height, and fresh weight over time.
  • Biomass Harvesting: During the logarithmic growth phase (for microalgae) or at maturity (for plants), harvest the biomass. Centrifuge microalgae cultures and wash the pellet. For plants, harvest the aerial parts.
  • Drying: Lyophilize or oven-dry the biomass to a constant weight to determine dry weight.
  • Biochemical Analysis:
    • Lipid Extraction: Use a solvent-based method (e.g., Bligh & Dyer) or Soxhlet extraction to isolate total lipids. Gravimetrically quantify lipid content as a percentage of dry weight [27].
    • Protein Quantification: Use the Bradford assay or Kjeldahl method to determine total protein content as a percentage of dry weight [28].
• Data Output: Biomass productivity (g/L/day), lipid content (% dry weight), protein content (% dry weight).

Experimental Workflow and Signaling Pathways

The experimental process for evaluating air revitalization efficiency involves a structured workflow from cultivation to data analysis. The following diagram visualizes the logical sequence and key decision points.

The Scientist's Toolkit: Research Reagent Solutions

The table below lists essential materials and reagents used in microalgae and plant research for air revitalization studies, along with their specific functions.

Reagent/Material	Function in Research	Example Use Case
Bold's Basal Medium (BBM)	A standardized nutrient medium providing essential macronutrients (N, P, K) and micronutrients for optimal microalgae growth [29] [30].	Used as a control medium to compare growth performance of different microalgae strains like Chlorella sorokiniana and Monoraphidium convolutum [30].
Tris-Acetate-Phosphate (TAP) Medium	A common mixotrophic/heterotrophic growth medium for microalgae; acetate provides a carbon source for growth in the dark [31].	Culturing model algae like Chlamydomonas reinhardtii for physiological and genetic studies [31].
Montmorillonite (Mt) Clay	A layered phyllosilicate used to study microalgae-mineral interactions, which can affect nutrient uptake, flocculation, and harvesting efficiency [31].	Investigating the biphasic effects of environmental particulates on algal physiology, such as growth and photosynthesis inhibition or enhancement [31].
Fluorescence Spectrophotometer	Instrument used to analyze extracellular polymeric substances (EPS) and photosynthetic pigments by measuring fluorescence signatures [31].	Characterizing the composition of EPS (proteins, polysaccharides) secreted by microalgae under different stress conditions [31].
Phyto-PAM-II Phytoplankton Analyzer	Measures chlorophyll fluorescence parameters (Fv/Fm, rETRmax) to assess the photosynthetic efficiency and health of microalgae [31].	Quantifying the inhibitory effect of stressors (e.g., clay minerals, pollutants) on the photosynthetic apparatus of microalgae [31].
Inductively Coupled Plasma Mass Spectrometry (ICP-MS)	A highly sensitive technique for quantifying elemental composition, including phosphorus uptake in algal cells [31].	Precisely measuring the phosphorus accumulation in microalgae biomass from the culture medium to study nutrient cycling [31].

From Lab to Application: Implementing Biological Systems for Air Revitalization

The escalating energy crisis and the urgent need for sustainable solutions have positioned microalgae as a cornerstone for green technologies. Unlike terrestrial plants, microalgae exhibit remarkably higher photosynthetic efficiency, enabling them to produce biomass up to ten times faster and more effectively [32]. This superior efficiency is particularly relevant for applications such as air revitalization in closed environments, where the continuous recycling of carbon dioxide and production of oxygen is paramount [33]. Photobioreactors (PBRs), which are closed systems designed for the phototrophic cultivation of microalgae, provide the controlled environment necessary to maximize these physiological advantages. This guide objectively compares the performance of predominant PBR configurations—Flat Panel, Bubble Column, Airlift, and Stirred Tank—by synthesizing experimental data on their hydrodynamic properties, mass transfer capabilities, and biomass productivity, with a specific focus on their implications for air revitalization efficiency.

Comparative Analysis of Photobioreactor Configurations

The design of a photobioreactor directly influences key parameters that dictate microalgal growth, including light penetration, gas transfer (CO₂ in and O₂ out), and mixing efficiency. The table below provides a systematic comparison of the most common closed PBR configurations.

Table 1: Performance Comparison of Major Photobioreactor Types for Microalgae Cultivation

Photobioreactor Type	Key Advantages	Key Limitations	Reported Biomass Productivity	Volumetric Mass Transfer Coefficient (kLa)	Suited for Air Revitalization
Flat Panel	High surface area-to-volume ratio; low oxygen buildup; high cell densities [34] [35]	Biofouling; can be difficult to scale up [34]	2.42 mg g⁻¹ (fucoxanthin yield) [34]	Data not available in search results	High (Efficient gas exchange and high biomass production) [34]
Bubble Column	Simple design; low cost; satisfactory heat and mass transfer [36] [37]	Low surface area-to-volume ratio can limit light harvesting [36]	0.097 gdw/L·day (C. sorokiniana) [36]	Highest among vertical columns [36]	Medium (Good mass transfer, but scaling can challenge light availability) [37]
Airlift	Low shear stress; defined fluid circulation; efficient mixing with low energy [34] [36]	Difficulty in scale-up; more complex design than bubble column [34]	0.072 gdw/L·day (C. sorokiniana) [36]	Can be limited, affecting growth [36]	High (Oriented flow through dark/light phases provides "flashing light effect") [36]
Stirred Tank	Optimal heat and mass transfer; high mixing efficiency [34] [36]	High shear stress can damage cells; high operating cost [34]	0.064 gdw/L·day (C. sorokiniana) [36]	Lower than bubble column, can limit growth [36]	Low (High shear and energy consumption are suboptimal for delicate cells) [34]

Experimental Protocols for PBR Performance Evaluation

To generate the comparative data presented in this guide, standardized experimental protocols are employed to assess the hydrodynamic and biological performance of different PBRs.

Protocol for Hydrodynamic and Mass Transfer Characterization

This methodology is critical for understanding the physical environment within a PBR, which directly impacts algal growth [36].

• Mixing Time (tₘ) Measurement: A tracer (e.g., acid or base) is injected into the reactor, and pH probes placed at various locations record the time required for the system to reach 95% of homogeneity. Lower mixing times (<10 seconds in characterized systems) indicate better homogenization, ensuring all cells have equal access to nutrients and light [36].
• Volumetric Mass Transfer Coefficient (kLa) Determination: The kLa, which quantifies the rate of oxygen transfer from the gas to the liquid phase, is measured using the gassing-out method. The dissolved oxygen (DO) concentration is first reduced by sparging with nitrogen. The reactor is then aerated, and the dynamic increase in DO is recorded. The kLa is calculated from the slope of a plot of ln(DO* - DO) versus time, where DO* is the saturation concentration. A higher kLa signifies more efficient oxygen removal and CO₂ dissolution [36].
• Gas Holdup (ε𝐺) Calculation: Gas holdup, the volume fraction of the reactor occupied by gas, is calculated as ε𝐺 = (Hf - H0) / Hf, where H₀ is the clear liquid height and H𝑓 is the aerated liquid height. Higher gas holdup generally correlates with better gas-liquid contact and mass transfer [36].

Protocol for Biological Performance Evaluation

This protocol assesses the direct outcome of PBR design on microalgal growth and compound production [34] [36].

• Strain and Cultivation: A target microalga (e.g., Phaeodactylum tricornutum for fucoxanthin or Chlorella sorokiniana for biomass) is inoculated into different PBRs containing a standardized growth medium like BG11 or F/2. Cultivation is typically performed in batch mode under controlled temperature and continuous illumination [34] [36].
• Growth Monitoring: Cell growth is tracked daily by measuring both cell density (using a haemocytometer or cell counter) and optical density (turbidity) at a specific wavelength, such as 680 nm [34].
• Biomass and Product Quantification: At the end of the cultivation period, biomass is harvested, and the dry cell weight is determined. For specific compounds like fucoxanthin, the biomass is subjected to extraction using solvents, and the target molecule is quantified via High-Performance Liquid Chromatography (HPLC) [34].

The Research Toolkit: Essential Reagents and Materials

Successful microalgal cultivation and experimentation rely on a suite of specific reagents and materials. The following table details key items used in the featured experiments.

Table 2: Essential Research Reagents and Materials for Microalgae Cultivation in PBRs

Item Name	Function/Application	Example Use Case
BG11 Medium	A defined nutrient solution providing essential macronutrients (Nitrogen, Phosphorus) and micronutrients (Iron, Boron) for freshwater microalgae growth [36] [38].	Cultivation of Chlorella sorokiniana and Chlorella vulgaris in bubble column and stirred tank PBRs [36] [38].
F/2 Medium	A widely used enriched seawater medium designed for the growth of marine microalgae and diatoms [34].	Cultivation of the diatom Phaeodactylum tricornutum for fucoxanthin production in flat panel PBRs [34].
Cationic Starch	A biodegradable, non-toxic flocculant used to aggregate microalgae cells into large flocs, significantly improving harvesting efficiency post-cultivation [39].	Pre-harvesting and concentration of Chlorella vulgaris biomass within an airlift PBR, enhancing final biomass concentration [39].
Computational Fluid Dynamics (CFD)	A powerful simulation tool used to model and optimize hydrodynamic parameters, fluid flows, and mass transfer within PBRs without building physical prototypes [34].	Simulating flow regime and mixing efficiency in a flat plate PBR to optimize its design for maximum fucoxanthin yield [34].

PBR Selection Workflow and Application Context

The choice of an optimal photobioreactor is a multi-faceted decision process that balances design principles with the end application. The following diagram maps out the logical pathway for selecting a PBR configuration, culminating in the specific context of air revitalization research.

Diagram: PBR Selection Workflow for Air Revitalization. The workflow begins with defining cultivation needs, evaluates key engineering criteria, and narrows down PBR options. For air revitalization, Flat Panel and Airlift PBRs are highly suitable due to superior gas exchange and controlled growth environments.

The selection of an appropriate photobioreactor is a decisive factor in harnessing the superior photosynthetic efficiency of microalgae for advanced applications like air revitalization. Experimental data confirms that no single PBR design is universally superior; each offers a distinct set of trade-offs. Flat panel PBRs excel in biomass and high-value product yield due to their high surface-to-volume ratio [34], while bubble column reactors offer an effective balance of performance and simplicity for scaled production [36] [37]. The controlled, low-shear environment of airlift PBRs is particularly well-suited for processes requiring high gas exchange and efficient mixing [39] [36]. When benchmarked against higher plants for air revitalization in closed systems, microalgae cultivated in optimized PBRs present a compelling advantage in terms of volumetric efficiency, metabolic versatility, and resilience, paving the way for their integration into next-generation life support and carbon capture systems.

Indoor air quality (IAQ) is a critical determinant of human health, well-being, and cognitive performance, with most people spending up to 90% of their time indoors [40] [41]. In the context of a broader thesis comparing microalgae and higher plants for air revitalization efficiency, this guide provides an objective comparison of two primary plant-based indoor air remediation technologies: traditional potted plants and advanced active green walls (AGWs). The pursuit of sustainable, biological solutions for maintaining air quality in closed environments, ranging from energy-efficient buildings to future space habitats, has accelerated research into these systems [42]. While potted plants represent a passive, nature-based approach, active green walls incorporate mechanical systems to enhance biofiltration, and emerging microalgae technologies promise even greater efficiency [43] [44]. This article synthesizes current experimental data to compare the performance, mechanisms, and applications of these systems, providing researchers and scientists with a clear, evidence-based guide.

Performance Comparison: Quantitative Data Analysis

The efficacy of plant-based systems in removing key airborne pollutants varies significantly based on their design, plant species, and operational mechanisms. The table below summarizes experimental data on the removal capabilities of potted plants, active green walls, and microalgae-based systems for common indoor pollutants.

Table 1: Air Pollutant Removal Performance of Different Biological Systems

System Type	Target Pollutant	Removal Performance	Experimental Conditions	Source
Potted Plants (Various Species)	CO₂	Reduction of 17-24% in sealed chambers; up to 51-77% when combined with ventilation	Sealed 1 m³ chamber; 24-hour exposure	[44]
Potted Plants (Golden Pothos)	CO₂	105 pots needed to absorb 208 ppm CO₂ in 80 min	Classroom with 13 students	[45]
Active Green Wall (Various Ferns)	Particulate Matter (PM)	45.78% (PM0.3–0.5) to 92.46% (PM5–10)	Controlled chamber study	[45]
Active Green Wall (Golden Pothos)	CO₂	Reduced concentration by ~46 ppm annually	Integrated with AC in a laboratory, annual study	[46]
Botanical Indoor Air Biofilter (BIAB) (Plants + Carbon Filter)	PM2.5 & VOCs	PM2.5: 5.36 µg/m³ per min; VOCs: 4.13 μg/m³ per min	Lab-scale biofilter rig with IoT sensors	[45]
Microalgae PBR (Spirulina maxima)	CO₂	~55% reduction from ~1100 ppm; up to ~90% from 10,000 ppm	0.064 m³ air chamber; NaHCO3-reduced medium	[44]

Beyond direct pollutant removal, these systems significantly impact the indoor environment. A year-long study on an Active Plant Wall (APW) showed it could bring mean skin temperature closer to the neutral 33.2°C and elevate perceptions of air freshness and thermal comfort to around “Fresh (+1)” and “Slightly comfortable (+1)” on subjective scales, demonstrating valuable psycho-physiological benefits [46].

Table 2: Additional Environmental and Functional Impacts

System Type	Thermal Regulation	Relative Humidity Impact	Psychological & Cognitive Benefits	Source
Active Green Wall	Decreased temp by 1.03°C - 1.35°C in winter/transition season	Increased RH by 11.6% - 20.76% in winter/transition season	Improved selective attention in children; enhanced perceived attention, creativity, and productivity in office workers	[46]
Potted Plants	Minor local effects	Can increase local humidity via evapotranspiration	Induced psychological relaxation and positive emotions; actual plants improved attention more than artificial ones	[41] [46]

Mechanisms and Experimental Protocols

Fundamental Removal Mechanisms

The air revitalization capabilities of these systems stem from biological and physical processes.

• Higher Plants (Potted Plants & AGWs): Plants primarily remove gases like CO₂ and VOCs through photosynthesis and phytoremediation [45]. The leaves absorb CO₂, while VOCs can be absorbed by the plant itself or metabolized by microorganisms in the rhizosphere (root zone) [45]. Particulate Matter (PM) is mainly removed through impaction and interception on the complex surface structures of leaves [41]. Furthermore, plants regulate the environment through transpiration, which releases water vapor and can increase humidity [46].
• Active Green Walls (AGWs): AGWs enhance these natural processes by using a fan to actively force air through the plant foliage and the growing substrate, which acts as a biofilter packed with pollutant-degrading microbes [45]. This active airflow dramatically increases the contact between polluted air and the biological components, leading to higher removal rates compared to passive potted plants.
• Microalgae Systems: In photobioreactors (PBRs), microalgae like Chlorella and Spirulina fix CO₂ through photosynthesis with an efficiency reported to be 10 to 50 times greater than that of terrestrial plants [43] [44]. They utilize pollutants as metabolic nutrients, releasing oxygen-rich, clean air as a byproduct.

Diagram 1: Air Revitalization Mechanisms Compared

Detailed Experimental Protocols

To ensure reproducibility and validate performance claims, researchers employ controlled experimental protocols. Key methodologies are detailed below.

Protocol for Testing Active Green Wall (AGW) Performance

This protocol is adapted from year-long studies assessing the impact of AGWs on indoor environmental quality [46].

• Objective: To quantitatively assess the impact of an Active Plant Wall (APW) on indoor air temperature, relative humidity, CO₂ concentration, and subjective human comfort across different seasons.
• Experimental Setup:
  • Chamber Configuration: Utilize two identical, adjacent laboratories. One serves as the experimental room (with APW), and the other as a control (without APW). Both rooms should have the same orientation, dimensions, and initial environmental conditions.
  • AGW System: The APW is integrated with the room's air-conditioning system. A common plant species like Epipremnum aureum (Golden Pothos) is cultivated in vertical planting slots with a sterile, well-draining growth medium (e.g., coconut husk, garden soil, vermiculite, perlite).
  • Sensor Placement: Install calibrated sensors to continuously monitor:
    • Air Temperature & Relative Humidity
    • CO₂ Concentration
    • Air Speed
  • Data Logging: Data should be collected automatically at frequent intervals (e.g., every 1-5 minutes) for at least one week per season (summer, transition season, winter) to capture temporal variations.
• Human Subject Evaluation:
  • Recruit participants to spend time in both the experimental and control rooms.
  • Measure participants' mean skin temperature (MST) using contact thermistors.
  • Administer standardized subjective questionnaires to assess perceptions of air freshness, thermal comfort, and overall well-being.
• Data Analysis:
  • Compare the time-averaged data (temperature, RH, CO₂) between the experimental and control rooms for each season.
  • Correlate objective environmental data with subjective human responses.
  • Perform statistical analysis (e.g., t-tests) to determine the significance of observed differences.

Diagram 2: AGW Testing Workflow

Protocol for Testing Microalgae Photobioreactor (PBR) Performance

This protocol is based on research investigating the CO₂ absorption performance of Spirulina maxima in a lab-scale PBR [44].

• Objective: To evaluate the growth and CO₂ sequestration efficiency of a microalgae species (Spirulina maxima) under a NaHCO₃-reduced cultivation medium in a controlled indoor environment.
• Experimental Setup:
  • Photobioreactor (PBR): A custom-designed, lab-scale PBR system with internal lighting and aeration capabilities.
  • Culture Medium: Prepare the standard SOT medium and a NaHCO₃-reduced amendment (e.g., a 1:1 v/v% mixture of standard and NaHCO₃-free medium).
  • Algae Inoculation: Inoculate Spirulina maxima into the PBR containing different medium amendments (standard, 50% NaHCO₃, NaHCO₃-free) to compare growth performance.
  • Test Chamber: Place the PBR inside a sealed air chamber of a known volume (e.g., 0.064 m³).
• Procedure:
  • Monitor algal cell growth (optical density, dry weight) for a prolonged period (e.g., 30 days).
  • Introduce a controlled concentration of CO₂ (~1100 ppm to simulate indoor air; 10,000 ppm for high-concentration tests) into the sealed chamber.
  • Use a CO₂ sensor to continuously monitor the concentration decay over time inside the chamber.
  • Record the time taken for the CO₂ level to saturate or reach a predetermined lower threshold.
• Data Analysis:
  • Calculate the CO₂ removal rate based on the reduction in concentration over time, normalized by the chamber volume and culture volume.
  • Compare the removal performance and algal growth kinetics across the different medium amendments.

The Scientist's Toolkit: Key Research Reagents & Materials

Successful experimentation in this field relies on a specific set of biological, chemical, and technological components.

Table 3: Essential Research Materials and Their Functions

Category	Item	Function in Research
Biological Components	Epipremnum aureum (Golden Pothos)	A model higher plant species due to its resilience, low light requirements, and known phytoremediation capabilities [45] [46].
	Spirulina sp. (e.g., S. maxima, S. platensis)	A model microalgae species valued for its rapid growth, high photosynthetic efficiency, and tolerance to environmental fluctuations [47] [44].
	Chlorella vulgaris	Another widely studied microalgae for CO₂ fixation and biomass production [47] [43].
Growth Substrates & Media	Sterile Nutrient Soil / Coconut Husk	The growth medium for higher plants in AGWs; provides physical support and hosts root microbiome essential for VOC degradation [46].
	SOT Medium / BBM	Standardized culture media for Spirulina and Chlorella, providing essential macro and micronutrients [44].
	Sodium Bicarbonate (NaHCO₃)	A primary carbon source in standard microalgae media; its reduction is studied to lower costs and simplify cultivation [44].
Monitoring & Analysis	RS485 Modbus Smart Sensors	Enable real-time, high-frequency monitoring of air quality parameters (PM, CO₂, VOCs, Temp, RH) in IoT-enabled experimental setups [45].
	Thermogravimetric Analysis (TGA)	A key technique for characterizing the thermal behavior and combustion properties of biomass waste from air purification systems [47].
	Fourier Transform Infrared (FTIR) Spectrometer	Coupled with TGA (TG-FTIR) to analyze gaseous products released during biomass pyrolysis, informing on energy recovery potential [47].

The experimental data clearly delineates the performance hierarchy and application niches for traditional and advanced plant-based systems. Potted plants offer a low-cost, passive solution with modest air purification benefits and valuable, well-documented psychological perks [41] [46]. However, their air revitalization capacity in real-world settings is limited, often requiring an impractically large number of plants to significantly impact air quality in a densely occupied space [40] [44].

Active Green Walls represent a significant technological evolution, enhancing natural biofiltration by actively forcing air through the plant-bed reactor. This results in quantifiably higher removal rates for CO₂, VOCs, and particulate matter, while also providing tangible thermal regulation and humidity control [45] [46]. The integration of IoT-based sensors allows for precise monitoring and control, making AGWs a robust option for improving Indoor Environmental Quality (IEQ) in modern buildings.

Emerging microalgae-based photobioreactors demonstrate a fundamentally different and highly efficient mechanism, particularly for CO₂ drawdown. Their performance, potentially 10-50 times greater than terrestrial plants, positions them as a promising solution for environments where compactness and high efficiency are paramount, such as in future Bioregenerative Life Support Systems (BLSS) for space exploration [43] [44] [42]. However, challenges related to the cost and complexity of cultivation medium management remain active areas of research [44].

In conclusion, the choice between these systems is application-dependent. Potted plants are suitable for settings where psychological benefits are the primary goal. For comprehensive air quality and environmental enhancement in commercial or institutional buildings, Active Green Walls are a proven, effective technology. For maximum CO₂ removal efficiency in closed-loop or highly polluted environments, microalgae PBRs represent the cutting edge of biological air revitalization research. Future work should focus on integrating these technologies—for example, combining AGWs with microalgae systems—to create synergistic, multi-tiered biofiltration systems for sustainable life support on Earth and beyond.

Integration with HVAC Systems and Building Infrastructure

The pursuit of improved indoor air quality and atmospheric revitalization within built environments has catalyzed research into biological air purification systems. Among these, two primary biological candidates emerge: terrestrial higher plants and photosynthetic microalgae. This guide objectively compares the integration potential and air revitalization performance of microalgae-based systems against those using higher plants within Heating, Ventilation, and Air Conditioning (HVAC) infrastructure. Framed within a broader thesis on air revitalization efficiency, this analysis provides researchers and scientists with experimental data, protocols, and technical considerations for evaluating these biogenic systems. The integration of biological components with mechanical building systems represents a frontier in sustainable building design, aiming to transform buildings from energy consumers into active, bioregenerative environments [48] [49].

Performance Comparison: Microalgae vs. Higher Plants

A quantitative comparison of microalgae and higher plants reveals significant differences in their core capabilities relevant to air revitalization. The following table summarizes key performance metrics derived from experimental observations and cultivation data.

Table 1: Performance Comparison of Microalgae and Higher Plants for Air Revitalization

Performance Metric	Microalgae Systems	Higher Plants (Terrestrial)
CO₂ Fixation Rate	1.6 - 2.0 g CO₂ / L culture / day [50]	3-6% of fossil fuel emissions annually [50]
CO₂ Fixation Efficiency	10 - 50 times higher than terrestrial plants [50] [7]	Baseline (1x)
Biomass Productivity	127 - 300 tons / hectare / year [50]	Varies significantly by species and climate
Primary Mechanism	Carbon Concentration Mechanism (CCM) [50]	C3 and C4 photosynthetic pathways [50]
Oxygen Production	Contributes ~50% of global O₂ [7]	Contributes ~50% of global O₂
Typical Integration Scale	Scalable Photobioreactors (PBRs) [50]	Green Walls, Indoor Potted Plants, Botanical Atria
Water Usage	Can utilize wastewater [7]	Typically requires potable water
Land Use Efficiency	High (vertical PBRs possible)	Lower (horizontal footprint typical)
Valuable Co-products	Biofuels, bioplastics, nutraceuticals [50] [7]	Limited in indoor settings

The data indicates that microalgae possess a superior CO₂ fixation rate and efficiency due to their fast growth and specialized Carbon Concentration Mechanism (CCM). This mechanism involves the pyrenoid organelle creating a CO₂-rich environment around the RuBisCO enzyme, drastically enhancing photosynthetic efficiency compared to the C3 and C4 pathways common in higher plants [50] [7]. Furthermore, microalgae systems offer higher scalability and a more direct pathway for carbon capture and conversion into valuable bio-products, making them particularly suited for integration with building infrastructure where space is at a premium [50].

Experimental Protocols for Performance Assessment

Protocol for Microalgae-CO₂ Fixation Kinetics

Objective: To quantify the CO₂ sequestration rate and biomass productivity of a microalgae strain under simulated HVAC-integrated conditions.

Materials:

• Photobioreactor (PBR): Glass or acrylic vessel with temperature, pH, and lighting control.
• Gas Mixing System: To simulate building flue gas or exhaust (e.g., 3-4% CO₂, air) [50].
• Analytical Instruments: CO₂ gas analyzer, spectrophotometer (for optical density), dry weight measurement setup.
• Cultivation Medium: Suitable growth medium (e.g., BG-11 for cyanobacteria).
• Microalgae Strain: Chlorella vulgaris or Spirulina (Arthrospira sp.), known for high CO₂ tolerance [50].

Methodology:

• Inoculation and Cultivation: Inoculate the sterile medium with the microalgae strain in the PBR.
• Parameter Control: Maintain constant temperature (e.g., 25-30°C), light intensity, and photoperiod. Continuously aerate the culture with the simulated exhaust gas at a controlled flow rate.
• Data Collection:
  • CO₂ Fixation: Measure the CO₂ concentration at the PBR inlet and outlet using the gas analyzer at 4-hour intervals. Calculate the CO₂ fixation rate using a mass balance approach [50].
  • Growth Monitoring: Record optical density (OD680) and dry cell weight daily to track biomass productivity.
• Analysis: Correlate CO₂ consumption with biomass increase. The fixation rate can be reported as g CO₂ / L culture / day.

Protocol for Higher Plant Phytoremediation Efficiency

Objective: To assess the removal rate of CO₂ and Volatile Organic Compounds (VOCs) by a higher plant species in a sealed chamber.

Materials:

• Sealed Test Chamber: Environmentally controlled with artificial lighting.
• Gas Analyzer: For CO₂ and specific VOCs (e.g., benzene, formaldehyde).
• Plant Species: Common indoor plants with purported air-purifying abilities (e.g., Spathiphyllum spp., Dracaena spp.).
• Soil and Potting System.

Methodology:

• Chamber Setup: Place a potted plant into the sealed chamber. A control chamber with only soil is recommended.
• Dosing and Monitoring: Inject a known quantity of CO₂ and/or VOCs into the chamber. Monitor the concentration decay over 24-48 hours using the gas analyzer.
• Environmental Control: Maintain constant light, temperature, and humidity throughout the experiment.
• Analysis: Calculate the removal rate based on the reduction in contaminant concentration over time, adjusted for the control chamber's data.

Protocol for HVAC-Integrated System Testing

Objective: To evaluate the performance impact of integrating a microalgae photobioreactor with a standard HVAC air handling unit (AHU).

Materials:

• Experimental HVAC System: A testbed AHU with standard components [51] [52].
• Pilot-Scale Photobioreactor: Designed for side-stream integration with the AHU return air.
• Sensor Network: Temperature, humidity, CO₂ sensors at AHU inlet, PBR inlet/outlet, and supply air [52].
• Energy Meter: To measure parasitic load from the PBR (pumps, lights).

Methodology:

• Baseline Measurement: Operate the AHU without the PBR integrated to establish baseline energy consumption and indoor CO₂ levels.
• Integrated Operation: Connect the PBR to treat a portion of the return air. The HVAC system supplies conditioned air to a simulated zone, with return air diverted through the PBR for CO₂ scrubbing and O₂ production [48].
• Data Collection: Record the change in CO₂ concentration across the PBR, the oxygenated air output, and the additional energy load imposed by the PBR system. Use standardized HVAC performance assessment methods for cross-domain fault diagnosis to ensure data reliability [51].
• Analysis: Calculate the net energy benefit by comparing the reduction in required outdoor ventilation air (and thus conditioning energy) against the energy cost of operating the PBR.

System Integration with Building Infrastructure

Integrating biological air revitalization systems requires careful consideration of building infrastructure and automation protocols. The workflow for integrating a microalgae photobioreactor with a smart building's HVAC system is illustrated below, highlighting the critical data and control exchanges.

(Diagram: HVAC-PBR Integration Workflow)

The successful implementation of this workflow relies on several technical pillars:

• Open Communication Protocols: Seamless integration is founded on open protocols like BACnet or Modbus, which enable interoperability between the HVAC equipment, the Building Automation System (BAS), and the photobioreactor's control system, regardless of manufacturer [48].
• Centralized Facility Management: Platforms like WebCTRL allow facility managers to monitor and control both the conventional HVAC and the integrated bioreactor from a single interface, providing centralized insight across systems [49].
• Data Leveraging for Decision-Making: The BAS uses real-time and historical data from both systems to enable predictive maintenance, optimize PBR operation based on occupancy patterns, and fine-tune the overall system for energy efficiency and air quality targets [53] [49].

The Scientist's Toolkit: Research Reagents & Essential Materials

For researchers designing experiments in this field, a core set of reagents and materials is essential. The following table details key items and their functions in experimental protocols.

Table 2: Essential Research Reagents and Materials for Air Revitalization Studies

Item	Function / Application
Chlorella vulgaris / Spirulina	Model microalgae organisms; known for high CO₂ tolerance and robust growth, making them ideal for foundational studies [50].
BG-11 Medium	A standard nutrient-rich cultivation medium optimized for the growth of cyanobacteria and some green algae.
CO₂ Gas Analyzer	Critical for quantifying the concentration of carbon dioxide at the inlet and outlet of test systems to calculate fixation/removal rates.
Photobioreactor (PBR)	A controlled bioreactor for cultivating phototrophic organisms with precise control over light, temperature, and gas exchange.
Spectrophotometer	Used to measure the optical density of microalgae cultures at 680nm (OD680), a proxy for biomass concentration.
Temperature & Humidity Chamber	Provides a stable, controlled environment for testing higher plant phytoremediation under reproducible conditions.
Building Automation System (BAS)	A centralized platform (e.g., WebCTRL) for monitoring and controlling integrated systems, essential for full-scale integration studies [49].
BACnet/IP Compatible Controller	Enables the experimental PBR or sensor network to communicate seamlessly with a standard building management system for data exchange and control [48].

The experimental data and integration analysis presented in this guide demonstrate a clear performance distinction between microalgae and higher plants for air revitalization. Microalgae systems exhibit a definitive advantage in CO₂ fixation rate, space efficiency, and co-product potential, making them a technically compelling candidate for deep integration with building HVAC infrastructure. Higher plants, while offering aesthetic and psychological benefits, function at a lower biochemical efficiency for targeted carbon capture.

The future of bioregenerative buildings lies in the seamless integration of these biological systems with smart building controls. Successful implementation requires a cross-disciplinary approach that combines microbiology, mechanical engineering, and data science. Researchers are encouraged to focus on optimizing photobioreactor energy consumption, developing robust control algorithms for dynamic building environments, and conducting long-term pilot studies to validate the durability and economic viability of these integrated systems.

As human space exploration ambitions extend beyond Low Earth Orbit to encompass Moon habitation and Mars missions, the development of self-sustaining life support systems has become increasingly critical. The current Environmental Control and Life Support System (ECLSS) on the International Space Station relies heavily on physicochemical processes that achieve approximately 85% water recovery and vent valuable methane byproducts into space, representing significant resource loss [11]. For long-duration missions where resupply is impractical, a more closed-loop, regenerative approach is essential.

Bioregenerative Life Support Systems (BLSS) offer a promising solution by using biological organisms to revitalize air, recycle water, and produce food. Within this context, a crucial scientific comparison emerges between microalgae (photosynthetic microorganisms) and higher plants (multicellular plants) for air revitalization efficiency. This review provides a structured, evidence-based comparison of these two biological systems, focusing on their respective capabilities in carbon dioxide absorption and oxygen production to support human life in isolated environments.

Performance Comparison: Microalgae vs. Higher Plants

Table 1: Comprehensive Performance Comparison of Microalgae and Higher Plants for Air Revitalization

Parameter	Microalgae	Higher Plants
Photosynthetic Efficiency	High (entire biomass photosynthetic)	Variable (non-photosynthetic structures present)
O2 Production Rate	Superior (faster growth and metabolic rates)	Moderate (slower growth and biomass accumulation)
CO2 Sequestration	Efficient, continuous	Diurnal pattern with reduced nighttime uptake
Space Requirements	Minimal (vertical cultivation possible)	Significant (horizontal footprint typically required)
Water Usage	Can utilize wastewater streams	Generally higher fresh water requirements
Harvesting & Processing	Requires specialized equipment	Simpler manual harvesting possible
Nutritional Co-product	High-value proteins, lipids, pigments	Direct food crops (fruits, vegetables)
System Startup Time	Days to weeks	Weeks to months
Resilience to Stress	High (rapid adaptation)	Variable (species-dependent)
Light Utilization	Efficient across spectrum	Specific wavelength requirements

Table 2: Quantitative Experimental Data from Air Purification Studies

Biomass Type	Treatment Scenario	Thermal Stability	Combustibility (HRC)	Key Findings
Chlorella vulgaris (Microalgae)	After CO2 absorption	High decomposition temperature	Significant heat release capacity	Compact structure post-treatment; high energy recovery potential
Arthrospira platensis (Microalgae)	After acetic acid exposure	Reduced thermal stability	Lower flammability	Structural degradation observed after pollutant exposure
Hedera helix (Higher Plant)	After toluene removal	Moderate thermal stability	Moderate combustibility	More compact and eroded surface post-treatment
Tillandsia xerographica (Higher Plant)	General air purification	High thermal resistance	High combustibility	Promising for energy valorization as combustion additive

Experimental Protocols for Efficiency Assessment

Photosynthetic Performance Measurement Protocol

Objective: Quantify photosynthetic and respiratory parameters of biological systems for air revitalization efficiency.

Materials:

• Photobioreactor or plant growth chamber
• Oxygen electrode system (e.g., Orion 97-08)
• CO2 analyzer (e.g., GC-7890B with TCD)
• Quantum meter (e.g., QMSS-SUN, Apogee Instruments)
• Temperature-controlled water bath
• Fresh biomass culture (microalgae or plant specimens)

Methodology:

• Sample Preparation: Culture microalgae or plants under standardized conditions. For microalgae, concentrate cells to 2-3 µg Chl mL⁻¹ via centrifugation. For plants, use leaf discs or whole seedlings [31] [54].
• System Setup: Transfer samples to water-jacketed gas exchange chamber supplemented with 10 mM NaHCO₃ to prevent carbon limitation.
• Parameter Measurement:
  • Photosynthesis: Measure O₂ evolution under increasing light intensities (0-2000 µmol photons m⁻² s⁻¹) using oxygen electrode.
  • Respiration: Measure O₂ consumption and CO₂ evolution in complete darkness.
  • Chlorophyll Fluorescence: Determine Fv/Fm (maximum quantum yield) and rETRmax (maximum relative electron transport rate) using pulsed amplitude fluorometry [31].
• Data Analysis: Calculate specific photosynthetic and respiratory rates normalized to biomass or chlorophyll content.

Key Calculations:

• Maximum specific growth rate (μmax) derived from log phase of growth curves
• Productivity: P = (Xᵢ - X₀)/tᵢ where X = biomass density
• Temperature coefficient (Q₁₀) for respiratory and photosynthetic processes [54]

Pollutant Removal Efficiency Protocol

Objective: Evaluate capacity of biological systems to remove specific air pollutants (CO₂, VOCs).

Materials:

• Sealed photobioreactor or plant growth chambers
• Gas chromatography system
• FTIR spectrometer
• Specific pollutant sensors (CO₂, toluene, acetic acid)
• Biomass samples (microalgae: Chlorella vulgaris, Arthrospira platensis; plants: Hedera helix, Tillandsia xerographica)

Methodology:

• System Setup: Place biomass in controlled environment chambers with continuous air mixing.
• Pollutant Introduction: Inject specific concentrations of target pollutants (CO₂: 400-5000 ppm; VOCs: 10-100 ppm).
• Monitoring: Track pollutant concentrations over time (0-72 hours) using integrated sensors and periodic GC/FTIR analysis.
• Post-Treatment Analysis:
  • Examine structural changes via SEM/EDX
  • Evaluate physiological changes through thermal analysis (TGA)
  • Assess metabolic changes via transcriptomic analysis [47] [31]
• Control Experiments: Run parallel systems without biomass for baseline comparison.

Metabolic Pathways in Air Revitalization

The metabolic pathways diagram illustrates the fundamental biological processes governing air revitalization in both microalgae and higher plants. While both systems utilize photosynthesis to convert CO₂ to O₂, microalgae employ enhanced CO₂ concentrating mechanisms that allow more efficient carbon fixation, particularly under elevated CO₂ conditions typical of spacecraft environments [11]. Higher plants regulate gas exchange through stomatal control, which provides response flexibility but can limit efficiency under suboptimal conditions.

Microalgae demonstrate faster metabolic turnover due to their simpler cellular organization and capacity for rapid cell division, translating to higher volumetric efficiency for O₂ production. Higher plants, however, distribute photosynthetic activity between specialized tissues, with significant carbon allocation to structural components that don't contribute directly to air revitalization [31].

Both systems can process volatile organic compounds (VOCs) through specialized metabolic pathways, though research indicates microalgae may have broader catalytic capabilities for diverse airborne contaminants [47].

The Scientist's Toolkit: Essential Research Reagents and Materials

Table 3: Essential Research Reagents and Equipment for Air Revitalization Studies

Category	Specific Items	Application Purpose	Key Features
Culture Systems	Photobioreactors, Growth chambers	Provide controlled environment for biomass cultivation	Temperature, light, and atmospheric control
Analytical Instruments	Oxygen electrode, CO2 GC-TCD, Phytoplankton PAM	Quantify gas exchange and photosynthetic parameters	Real-time monitoring capability
Microalgae Strains	Chlorella vulgaris, Arthrospira platensis	Primary photosynthetic organisms for study	High photosynthetic efficiency, well-characterized
Higher Plant Species	Hedera helix, Tillandsia xerographica	Comparative plant systems for air revitalization	Known air purification capabilities
Growth Media	TAP medium, Basal nutrient medium	Provide essential nutrients for growth	Standardized composition for reproducibility
Pollutant Sources	CO2 tanks, VOC standards (toluene, acetic acid)	Simulate contaminated air environments	Precise concentration control
Molecular Biology Tools	RNA-Seq kits, SEM-EDX equipment	Analyze physiological and genetic responses	Reveal mechanism of action at cellular level
Thermal Analysis	TGA, Microscale Combustion Calorimetry	Assess biomass properties and energy content	Evaluate byproduct valorization potential

Comparative Analysis: Advantages and Limitations

Microalgae Systems

Advantages:

• Higher Photosynthetic Efficiency: Microalgae achieve superior O₂ production per unit volume due to their simple cellular structure and entire biomass being photosynthetic-active [11].
• Continuous Operation: Unlike higher plants that exhibit reduced photosynthetic activity during dark periods, microalgae systems can be operated with continuous illumination for constant air revitalization.
• Waste Integration Capability: Microalgae can utilize wastewater streams (e.g., urine processing assembly outputs) as nutrient sources, enhancing resource closure in BLSS [11].
• Adaptive Metabolism: Studies demonstrate that microalgae like Chlorella sp. can rapidly acclimate to varying temperature and light conditions, maintaining performance under environmental fluctuations [54].

Limitations:

• Downstream Processing Requirements: Harvesting microalgae biomass from liquid cultures requires specialized equipment such as centrifuges or filtration systems.
• Cultural Acceptability: As a food source, microalgae may face resistance due to unfamiliarity, though they offer high-quality proteins and nutrients [11].
• System Stability: Monocultures of microalgae may be vulnerable to contamination or crash without careful monitoring and control.

Higher Plant Systems

Advantages:

• Direct Food Production: Systems like lettuce cultivation provide familiar food crops along with air revitalization, with studies showing yield increases of up to 18.3% with biostimulant applications [29].
• Psychological Benefits: The presence of higher plants may offer psychological advantages for crew members during long-duration missions.
• Structural Diversity: Complex architecture provides varied microenvironments that could support other biological processes.
• Cultural Familiarity: Well-established cultivation protocols and immediate recognition as food sources.

Limitations:

• Slower Response Times: Longer generation times and physiological inertia limit rapid adaptation to changing atmospheric conditions.
• Resource Intensive: Generally require greater mass, volume, and water resources per unit O₂ production compared to microalgae systems.
• Diurnal Variation: Photosynthetic activity follows light-dark cycles, creating periodic fluctuations in air revitalization capacity.

The comprehensive analysis of experimental data and case studies presented herein demonstrates that both microalgae and higher plants offer distinct advantages for air revitalization in controlled environments. Microalgae systems generally provide superior performance in terms of volumetric efficiency and adaptation capability, making them particularly suitable for space-constrained applications where rapid response to atmospheric changes is critical. Conversely, higher plant systems offer the dual benefit of familiar food production alongside air revitalization, with potential psychological advantages for crew members.

Future research should focus on integrated approaches that leverage the strengths of both biological systems. Specifically, investigation into multi-trophic systems combining rapid-response microalgae-based air revitalization with higher plants for food production and psychological benefits could optimize overall BLSS performance. Additionally, advances in genetic engineering of microalgae, particularly for enhanced CO₂ fixation pathways and stress resistance, present promising avenues for significantly improving system efficiency and reliability [55].

The increasing availability of automated phenotyping platforms, such as the PhenoSelect system that enables high-throughput screening of microalgal strains under multiple environmental conditions, will accelerate the identification and development of optimal strains for air revitalization applications [56]. Furthermore, exploration of psychrophilic microalgae species, which exhibit unique adaptation mechanisms to extreme environments, may yield valuable biological components for BLSS requiring operation under variable temperature conditions [54].

As human space exploration continues to evolve, the integration of robust biological systems for life support will be essential for mission success. The experimental protocols, performance data, and comparative analysis provided in this review serve as a foundation for informed decision-making in the design of next-generation air revitalization systems.

Overcoming Limitations: Strategies for Enhancing Efficiency and Scalability

Addressing Light Utilization Inefficiencies and Photoinhibition

The efficiency of photosynthetic air revitalization—the process of converting carbon dioxide (CO₂) to oxygen (O₂) in closed-loop systems—is fundamentally constrained by how effectively photosynthetic organisms capture and utilize light energy. Both microalgae and higher plants are primary producers capable of this conversion; however, their light-harvesting systems face inherent inefficiencies. A major limitation arises from the disproportionality between the fast rate of photon capture by light-harvesting antennae and the slower rate of downstream photosynthetic electron transfer, leading to energy losses that can exceed 50% at full sunlight [57]. When light intensity exceeds the photosynthetic capacity, it leads to photoinhibition, a decline in the efficiency of Photosystem II (PSII), and the overproduction of harmful reactive oxygen species (ROS) [58] [59] [60]. This comparative guide analyzes the distinct photoprotective strategies and experimental data for microalgae and higher plants, providing a framework for selecting and optimizing organisms for efficient air revitalization systems.

Comparative Analysis of Photoprotective Mechanisms

Photosynthetic organisms have evolved a suite of mechanisms to mitigate light stress. The strategies employed by microalgae and higher plants share common principles but differ in their emphasis and regulation.

Table 1: Comparison of Photoprotective Mechanisms in Microalgae and Higher Plants

Mechanism	Microalgae	Higher Plants	Function in Photoinhibition Prevention
Non-Photochemical Quenching (NPQ)	Rapid, zeaxanthin-dependent; enhanced under high light and nutrient stress [61] [62].	Multi-component (qE, qZ, qI); involves PsbS protein and xanthophyll cycle [63] [60].	Dissipates excess light energy as heat, preventing ROS generation.
Antioxidant Systems	Synthesis of antioxidant carotenoids (e.g., β-carotene, astaxanthin, fucoxanthin) [59] [62].	Enzymatic scavengers (e.g., CAT, SOD); non-enzymatic antioxidants like anthocyanins [60] [64].	Scavenges reactive oxygen species (ROS) produced under light stress.
Light-Harvesting Antenna Tuning	Reduction in chlorophyll b content tunes antenna size, boosting photosynthetic rate under high light [57].	Increased chlorophyll a/b ratio and reduced antenna size under acclimation [58] [60].	Reduces the optical cross-section, minimizing over-excitation.
Chloroplast Movement	Not typically applicable in unicellular species.	Avoidance response to reposition chloroplasts under high light [60].	Minimizes light absorption by moving organelles away from direct light.
Photorespiration	Possess CO₂ Concentrating Mechanisms (CCMs) to suppress photorespiration [62].	Key pathway for ROS metabolism; essential for high light tolerance (e.g., HPR1 role) [64].	Consumes excess energy and metabolites, maintaining redox balance.

Fundamental Differences in Strategic Approach

The data reveals a fundamental divergence in strategy. Microalgae heavily rely on biochemical photoprotection through the rapid modulation of NPQ and the accumulation of secondary carotenoids, which also represent valuable bioproducts [65] [59]. Their CCMs make them highly efficient at carbon fixation under a range of conditions, directly suppressing the photorespiration pathway that is a major source of ROS in higher plants [62]. In contrast, higher plants employ more structural and metabolic adaptations, including complex chloroplast movements and a tightly integrated photorespiratory pathway that is essential for managing oxidative stress under high light [60] [64]. The HPR1 enzyme in Arabidopsis, for instance, is critical for maintaining the dynamic balance of ROS and photorespiration under high light stress [64].

Quantitative Performance Data under Light Stress

The physiological impact of light stress is quantifiable through key photosynthetic parameters. The following table summarizes experimental data from various studies.

Table 2: Quantitative Performance Metrics under Different Light Conditions

Organism	Light Condition	Key Impact on Growth	Impact on Photosynthesis (Fv/Fm)	Pigment / Metabolite Response
Chlorella sorokiniana [65]	Violet light, high CO₂	Max. growth: 5.89 log10 cells/mL	Not Specified	Max. chlorophyll: 0.3049 µg/mL
Chlorella vulgaris [59]	26-400 µmol m⁻² s⁻¹ (Optimal)	Optimal growth range	Not Specified	Increased lipid synthesis at >60 µmol m⁻² s⁻¹
Sargassum fusiforme [61]	300 vs. 30 µmol m⁻² s⁻¹	Not Specified	Fv/Fm decreased	XCP/Chl a & Fx/Chl a ratios increased
Arabidopsis thaliana [58]	Periodic High Light (1800 µmol m⁻² s⁻¹)	Lower leaf area, higher seed yield	Initial photoinhibition, then acclimation	Chlorophyll a/b ratio increased
Arabidopsis hpr1 mutant [64]	High Light (350 µmol m⁻² s⁻¹)	Severe growth retardation	Serious photoinhibition	Excessive ROS, high photorespiratory intermediates

Analysis of Comparative Data

The data demonstrates that both systems can acclimate to high light, but the outcomes differ. Microalgae like Chlorella can achieve high biomass and pigment productivity under specific, optimized light spectra [65]. Furthermore, elevated light intensity is a reliable trigger for increasing the yield of valuable metabolites such as lipids and carotenoids [59]. In higher plants, Arabidopsis shows remarkable resilience through acclimation, recovering from initial photoinhibition and even increasing reproductive yield despite reduced leaf area [58]. However, the failure of the hpr1 mutant to thrive under high light underscores the critical, non-redundant role of an efficient photorespiratory pathway in plants for managing light stress [64].

Essential Experimental Protocols for Analysis

To evaluate light utilization efficiency and photoinhibition, researchers employ a standardized set of protocols. Below are detailed methodologies for key assays.

Chlorophyll Fluorescence Measurement (PAM Fluorometry)

This is a non-invasive, rapid technique to assess the photochemical efficiency of PSII.

• Sample Preparation: Dark-acclimate the leaf or algal culture for at least 30 minutes to ensure all reaction centers are open and relax energy-dependent quenching [61] [63].
• Initial Fluorescence Measurement: Apply a low-intensity measuring beam to determine the initial fluorescence level, ( F_o ).
• Maximal Fluorescence Measurement (( Fm )): Apply a saturating pulse of light (e.g., 0.2-1.0 s, >3000 µmol m⁻² s⁻¹) to close all PSII reaction centers. The recorded fluorescence peak is ( Fm ) [63].
• Calculation of Maximal PSII Efficiency: Compute the variable to maximal fluorescence ratio, ( Fv/Fm = (Fm - Fo)/F_m ). A value of ~0.83 is typical for healthy, unstressed plants and algae. A decrease indicates photoinhibition [58] [61].
• NPQ Induction: Expose the sample to actinic light (e.g., 1150 µmol m⁻² s⁻¹ for 5 minutes) while periodically applying saturating pulses. NPQ is calculated as ( (Fm - Fm')/Fm' ), where ( Fm' ) is the maximal fluorescence under light [61].

Photosynthetic Oxygen Evolution Measurement

This method directly measures the rate of photosynthesis as O₂ production.

• Apparatus Setup: Use an oxygen electrode chamber (e.g., Hansatech) maintained at a constant temperature (e.g., 25°C) [63].
• Sample Loading: For leaves, use a leaf disc. For microalgae, concentrate a known volume of culture and introduce it into the chamber.
• Illumination and Data Recording: Illuminate the sample with a range of light intensities (Photosynthetically Active Radiation, 400-700 nm). Record the rate of O₂ production once it stabilizes at each intensity [63].
• Data Analysis: Plot the rate of O₂ evolution against light intensity to generate a light response curve, determining the light-saturated rate of photosynthesis and the quantum yield of O₂ evolution in limiting light [63].

Pigment Quantification via HPLC

This protocol provides precise quantification of photosynthetic and photoprotective pigments.

• Pigment Extraction: Homogenize frozen leaf or algal pellet in 100% methanol, dimethylformamide, or acetone. Centrifuge to pellet debris [65] [61].
• HPLC Analysis: Inject the supernatant into a High-Performance Liquid Chromatography (HPLC) system equipped with a C18 reverse-phase column.
• Separation and Detection: Use a gradient elution with a mixture of solvents (e.g., acetonitrile, methanol, ethyl acetate) to separate pigments. Detect pigments using a photodiode array detector by their specific absorbance spectra (e.g., Chl a at 430 and 664 nm, carotenoids at 450 nm) [61].
• Quantification: Calculate pigment concentrations by comparing peak areas to those of known standards. Results are often expressed as ratios to chlorophyll a (e.g., XCP/Chl a, Fx/Chl a) to indicate relative pool sizes [61].

Diagram 1: Divergent photoprotection signaling pathways in microalgae and higher plants. While both respond to overexcitation and ROS, their strategic priorities differ, leading to distinct physiological outcomes.

The Scientist's Toolkit: Key Research Reagent Solutions

Table 3: Essential Reagents and Tools for Photosynthesis Stress Research

Reagent / Tool	Function / Application	Example Use Case
PAM Fluorometer	Measures chlorophyll fluorescence parameters (Fv/Fm, NPQ) in vivo.	Quantifying the extent of photoinhibition and capacity for photoprotection in intact leaves/algae [58] [61].
Oxygen Electrode	Directly measures the rate of photosynthetic O₂ evolution or consumption.	Generating light response curves and quantifying the quantum yield of photosynthesis [63].
LED Light Sources	Provides precise control over light intensity, quality (spectrum), and photoperiod.	Studying the effect of specific wavelengths (e.g., violet vs. red) on growth and pigment production in Chlorella [65] [59].
HPLC System with PDA Detector	Separates, identifies, and quantifies individual pigments and metabolites.	Precisely measuring the ratios of xanthophyll cycle pigments (Vx, Ax, Zx) to Chl a in Sargassum [61].
Photorespiratory Mutants	Genetic models (e.g., hpr1) to dissect the role of specific pathways in stress response.	Elucidating the essential role of HPR1 in high light tolerance in Arabidopsis [64].
Artificial Seawater & PESI	Controlled nutrient media for algal cultivation, allowing nutrient stress studies.	Investigating the combined effects of irradiance and nutrient availability on NPQ in Sargassum fusiforme [61].

Diagram 2: A standard experimental workflow for evaluating light stress responses, integrating physiological and biochemical assays.

The comparative analysis reveals that the choice between microalgae and higher plants for air revitalization is context-dependent. Microalgae demonstrate superior potential for rapid biomass production and intrinsic integration of high-light stress with valuable metabolite synthesis, making them ideal for bioreactor-based systems where volume and speed are critical. Their CCMs provide a distinct advantage in efficient carbon fixation. Conversely, higher plants exhibit more complex, whole-organism acclimation strategies that prioritize long-term survival and reproductive success, which could be advantageous for multi-purpose life support systems providing food and psychological benefits.

Future research should focus on leveraging the strengths of each system through emerging technologies. For microalgae, metabolic engineering to further optimize light-harvesting antenna size and enhance carotenoid pathways is promising [57] [62]. For higher plants, breeding or engineering varieties with enhanced photoprotective capacity, such as faster NPQ relaxation or altered photorespiratory flux, could boost overall efficiency [60] [64]. The decision framework ultimately hinges on the specific mission parameters: microalgae for maximal O₂ production and CO₂ sequestration per volume, and higher plants for multi-functional, resilient ecological systems.

Nutrient Management and Strain Selection for Robust Performance

The challenge of maintaining breathable air in closed environments, from spacecraft to advanced terrestrial facilities, has intensified research into biological air revitalization systems. This domain pits two primary biological contenders against each other: traditional higher plants and microscopic algae. While higher plants provide psychological benefits and food co-products, microalgae possess distinct physiological advantages for efficient gas exchange. Microalgae exhibit photosynthetic efficiency far surpassing terrestrial plants, enabling them to capture CO2 and generate O2 at significantly higher rates per unit volume [66]. Their rapid growth cycles and ability to thrive in controlled bioreactors without soil make them exceptionally suited for integration into engineered life support systems [67]. This guide provides an objective comparison of performance outcomes based on different nutrient management strategies and strain selection, delivering critical data for researchers and scientists optimizing these systems for applied use.

Performance Comparison of Microalgae Strains and Cultivation Systems

The effectiveness of a microalgae-based air revitalization system is fundamentally governed by the choice of species and the cultivation conditions. The table below summarizes experimental data for several promising strains.

Table 1: Performance Comparison of Selected Microalgae Strains for Air Revitalization

Microalgae Strain	Max Biomass Yield (g/L)	Optimal Temp (°C)	Optimal pH	CO2 Capture Rate	Key Biomolecules	Tolerance to Co-culture
Chlorella vulgaris	0.83 [67]	30 [67]	4.0-8.0 [67]	High [66]	Proteins, Lipids [67]	High (with bacteria) [68]
Scenedesmus sp.	0.80 [67]	37 (thermotolerant) [69]	~7.0 [67]	High [66]	Lipids, Carbohydrates [67]	High (with mammalian cells) [69]
Chlorella sorokiniana	N/A	37 (thermotolerant) [69]	~7.0	Moderate to High [66]	Lipids, Pigments [67]	Moderate [69]
Chlamydomonas reinhardtii	0.79 [67]	30 [67]	6.0-8.0 [67]	Moderate [66]	Lipids, Hydrogen [70]	Low to Moderate [69]
Nannochloropsis gaditana	0.73 [67]	30 [67]	~7.0 [67]	High [66]	PUFAs, Pigments [67]	N/A

The cultivation system design directly impacts biomass productivity and, consequently, gas exchange performance. The following table compares two primary reactor types.

Table 2: Comparison of Microalgae Cultivation System Configurations

Parameter	Open Ponds (OPs)	Photobioreactors (PBRs)
Volumetric Productivity	Lower [66]	Higher [66]
CO2 Capture Efficiency	Lower (subject to atmospheric loss) [66]	Higher (controlled environment) [66]
Resource Control (Nutrients, pH, Temp)	Difficult [66]	Precise [66]
Risk of Contamination	High [68]	Low [68]
Land/Area Requirement	High [66]	Lower (vertical stacking possible) [66]
Capital & Operational Cost	Lower	Higher
Suitable Strains	Robust, fast-growing (e.g., Chlorella, Scenedesmus) [67]	High-value, sensitive strains [66]

Experimental Protocols for Performance Evaluation

Protocol 1: Optimizing Symbiotic Microalgae-Bacteria Co-culture

Objective: To determine the optimal cultivation parameters for a symbiotic microalgae-bacteria system to maximize biomass and lipid productivity for integrated wastewater treatment and bioenergy feedstock production [68].

Materials:

• Strains: Indigenous wastewater-borne microalgae and bacteria, separated via multi-step filtration (5 μm for microalgae, 0.2 μm for bacteria) [68].
• Medium: Synthetic wastewater (e.g., 0.4125 g/L glucose, 0.078 g/L NH₄Cl, 0.018 g/L KH₂PO₄, trace metals) [68].
• Equipment: Baffled Erlenmeyer flasks, table top shaker, flow cytometer (e.g., BD Accuri C6 Plus), CO₂-air mixing system, adjustable light source [68].

Methodology:

• Experimental Design: Employ a Central Composite Design (CCD) with five levels for each of the four factors:
  • Inoculum ratio of bacteria to microalgae (25% to 100%)
  • CO₂ supply (1% to 5% v/v)
  • Light intensity (50 to 300 μmol/m²/s)
  • Harvest time (1 to 15 days) This generates 30 experimental runs [68].
• Cultivation: Inoculate cultures in 500 mL baffled flasks with constant shaking (100 RPM) under a 12-h light/12-h dark cycle. Maintain CO₂ flow at 0.25 vvm [68].
• Monitoring & Analysis: Harvest cultures according to the design matrix. Analyze microalgal biomass concentration (cells/mL) and lipid productivity (Total Fluorescent Units/mL) using flow cytometry [68].
• Optimization: Use Response Surface Methodology (RSM) to fit quadratic models to the experimental data. Determine the optimal combination of factors that maximizes both biomass and lipid productivity [68].

Protocol 2: High-Throughput Selection of Strains via Competitive Phototaxis

Objective: To rapidly isolate microalgal strains with superior photosynthetic efficiency from large mutant libraries using a microfluidic device that selects for fast phototactic response [70].

Materials:

• Strains: Wild-type (e.g., Chlamydomonas reinhardtii CC-125) and a library of its random insertional mutants (>10,000) [70].
• Equipment: Poly(dimethylsiloxane) (PDMS) microfluidic device with a long, gradually narrowing channel; green LED light source (70 μmol photons m⁻²s⁻¹); high-speed camera [70].
• Reagents: TAP or other standard culture medium [70].

Methodology:

• Device Preparation: Load the microalgal cell mixture into the inlet of the microfluidic channel [70].
• Phototaxis Assay: Expose the channel to a directional green LED light source. Motile cells will exhibit negative phototaxis, swimming away from the light source toward the outlet [70].
• Cell Sorting & Analysis: Monitor and record the arrival time of individual cells at a designated observation zone near the outlet using a high-speed camera. Isolate the cells that arrive first, representing those with the fastest phototactic response [70].
• Validation: Culture the isolated strains and quantitatively evaluate their photosynthetic performance by measuring photosystem II operating efficiency (Y(II)) and photoautotrophic growth rate compared to the parental strain [70].

Protocol 3: Model-Predictive Nutrient Control Under Light/Dark Cycles

Objective: To optimize glucose and nitrate feeding during alternating light and dark cycles to maximize biomass yield and nutrient utilization efficiency using genome-scale metabolic models (GSMs) [71].

Materials:

• Strain: Chlorella vulgaris [71].
• Medium: Standard culture medium with variable glucose and nitrate [71].
• Equipment: Photobioreactor, OD₇₅₀ spectrophotometer, nutrient analyzers [71].
• Software: Genome-scale metabolic models for C. vulgaris (iCZPA-T1 for photoautotrophic growth, iCZH-T1 for heterotrophic growth) [71].

Methodology:

• System Setup: Cultivate C. vulgaris in a bioreactor under alternating 16-hour light and 8-hour dark cycles [71].
• Monitoring: Measure biomass (OD₇₅₀) and residual glucose/nitrate concentrations every 8 or 16 hours [71].
• Model-Predictive Feeding:
  • During Dark Cycle: Apply the heterotrophic model (iCZH-T1), constrained by current biomass, to predict the precise amount of glucose and nitrate required to support growth for the next 8 hours. Add the predicted amounts at the start of the dark cycle [71].
  • During Light Cycle: Switch to the photoautotrophic model (iCZPA-T1) to predict nitrate requirements for light-driven growth, adding the predicted amount while CO₂ is supplied [71].
• Performance Analysis: Compare final biomass density, lutein, and fatty acid yields per gram of consumed glucose against control cultures grown in standard heterotrophic or autotrophic batch modes [71].

Visualization of Workflows and Metabolic Pathways

High-Throughput Strain Selection Workflow

This diagram illustrates the integrated protocol for selecting high-performance microalgae strains using microfluidic phototaxis screening.

Model-Predictive Nutrient Control Logic

This diagram outlines the closed-loop control system for precision nutrient feeding based on genome-scale metabolic models.

The Scientist's Toolkit: Essential Research Reagents and Materials

The following table details key reagents and materials essential for conducting research in microalgae nutrient management and strain selection.

Table 3: Key Research Reagent Solutions for Microalgae Cultivation and Analysis

Reagent/Material	Function/Application	Example Usage in Protocols
Synthetic Wastewater Medium	Provides standardized, controllable nutrient base for experimentation without the variability of real wastewater.	Serves as the growth medium in co-culture optimization studies [68].
Alginate-Based Hydrogel	A biocompatible polymer for immobilizing microalgae in 3D bioprinted constructs for co-culture studies.	Used to embed Scenedesmus sp. for photosynthetic oxygenation in mammalian cell co-cultures [69].
Genome-Scale Metabolic Models (GSMs)	In silico models of microalgal metabolism used to predict nutrient demands and optimize feeding strategies.	iCZPA-T1 and iCZH-T1 models for Chlorella vulgaris predict nitrate/glucose needs in light/dark cycles [71].
Microfluidic Phototaxis Device (PDMS)	A high-throughput platform for analyzing and selecting microalgal cells based on their behavioral response to light.	Enables isolation of C. reinhardtii mutants with fast phototaxis and high photosynthetic efficiency [70].
Flow Cytometer with Fluorescence Detection	Enables rapid quantification of microalgal biomass, cell size, and neutral lipid content at a cellular level.	Used for high-throughput analysis of biomass and lipid productivity in RSM experiments [68].

Genetic and Metabolic Engineering to Boost Photosynthetic Rates and Pollutant Uptake

The escalating challenges of climate change and environmental pollution have intensified the search for sustainable biological solutions. Within this context, enhancing the photosynthetic efficiency and pollutant uptake capabilities of photosynthetic organisms presents a transformative opportunity. This guide provides a comparative analysis of two principal groups—microalgae and higher plants—evaluating their potential as platforms for genetic and metabolic engineering, with a specific focus on applications for air revitalization. Microalgae, comprising diverse eukaryotic protists and prokaryotic cyanobacteria, exhibit remarkable physiological adaptability and high photosynthetic efficiency [72] [7]. In contrast, higher plants (C3 species) offer the advantage of complex development and direct integration into agricultural ecosystems but are hampered by the inherent inefficiencies of the Rubisco enzyme [73] [74]. This article objectively compares the performance of engineered microalgae and higher plants, drawing on experimental data to delineate their respective advantages, limitations, and future prospects.

Performance Comparison: Engineered Microalgae vs. Higher Plants

The strategic application of genetic engineering has targeted key biological processes in both microalgae and higher plants. The tables below synthesize experimental data related to the enhancement of photosynthetic efficiency and pollutant removal capabilities.

Table 1: Comparative Performance in Photosynthetic Enhancement

Organism/Strategy	Genetic Modification	Key Experimental Findings	Theoretical/Measured Gain
Microalgae (General)	Native Carbon Concentrating Mechanism (CCM)	Concentrates CO₂ around Rubisco via pyrenoid [75] [74]	Accounts for ~50% of global C fixation [74]
Higher Plant (Arabidopsis)	Introduction of algal CCM components (Rubisco + EPYC1)	Formation of proto-pyrenoid condensates in chloroplasts [74]	Model predicts up to 60% increase in photosynthetic efficiency [74]
Higher Plant (Theoretical)	Replacement of RuBisCO with PEPC via MOG cycle	Proposed metabolic bypass for RuBisCO's limitations [73]	Potential for significant increase in efficiency [73]
Higher Plant (Tobacco)	Overexpression of Cytochrome b6f Rieske FeS protein	Increased electron transport rates and growth under high light [76]	Increased photosynthetic efficiency and growth [76]
Higher Plant (Tobacco)	Overexpression of Ferredoxin-NADP+ reductase (FNR)	Increased NADPH availability for Calvin cycle [76]	Increased photosynthetic capacity and growth [76]

Table 2: Comparative Performance in Pollutant Uptake and Bioremediation

Organism	Target Pollutant	Experimental Findings & Performance	Engineering Approach
C. vulgaris	Nitrogen, Phosphorus, COD (Wastewater)	80-94% removal of total N and P; 72% COD removal [72]	Use of wild-type and engineered strains; robust species [72]
C. vulgaris	Heavy Metals (e.g., Ca, Mg, Mo)	99%, 85%, and 42% removal of Ca, Mg, and Mo, respectively [72]	Natural bioaccumulation; potential for enhanced metal-binding peptides [72]
P. tricornutum	Nitrate, Phosphate, Iron (Oilfield Water)	92%, 76%, and 85% removal of nitrate, phosphate, and iron [72]	Cultivation in saline wastewater; genetic toolkits available [72]
Engineered Microalgae	Pesticides (Atrazine, Carbofuran, etc.)	96–99% removal efficiency of common pesticides [72]	Expression of specific pesticide-degrading enzymes [72]
Engineered Microalgae	Plastics (e.g., PET)	Creation of PET plastic-eating microalgae [72]	Heterologous expression of bacterial PET-degrading enzymes [72]
Higher Plants	Heavy Metals, VOCs (Air/Soil)	Moderate efficiency; limited by growth rate and biomass [76] [7]	Overexpression of aquaporins, metal transporters, and TF genes [76]

Detailed Experimental Protocols

To facilitate replication and further research, this section outlines detailed methodologies for key experiments cited in the performance comparison.

Protocol: Engineering a Proto-Pyrenoid in a Higher Plant

This protocol is based on the groundbreaking work of engineering a carbon-concentrating mechanism (CCM) into Arabidopsis thaliana [74].

Objective: To form a liquid-like Rubisco condensate (proto-pyrenoid) in the chloroplast of a C3 plant to enhance CO₂ concentration around the enzyme.

Key Reagents:

• Plant Material: Arabidopsis thaliana wild-type and mutant lines.
• Genetic Constructs:
  • pEA1: A plant transformation vector containing a gene encoding a hybrid plant-algal Rubisco large subunit (from tobacco engineered with Chlamydomonas Rubisco sequences) to enable binding to the linker protein EPYC1.
  • pEA2: A plant transformation vector containing the gene for the algal linker protein EPYC1 (Essential Pyrenoid Component 1), fused to an Arabidopsis chloroplast transit peptide to ensure localization to the chloroplast.
• Culture Media: Standard MS (Murashige and Skoog) agar plates and liquid medium for plant growth and selection.
• Selection Agents: Appropriate antibiotics (e.g., Kanamycin, Hygromycin) depending on the selectable marker genes used in the constructs.

Methodology:

• Plant Transformation: Introduce the constructed plasmids (pEA1 and pEA2) into Arabidopsis plants using the floral dip method with Agrobacterium tumefaciens strain GV3101.
• Selection and Growth: Select transformed seeds (T1 generation) on MS plates containing antibiotics. Grow resistant seedlings under controlled environmental conditions (22°C, 16/8h light/dark cycle).
• Genotyping: Confirm the integration and homozygosity of the transgenes in subsequent generations (T2, T3) using PCR and Southern blot analysis.
• Protein Expression Analysis: Verify the expression and chloroplast localization of the hybrid Rubisco and EPYC1 proteins via Western blotting and immunocytochemistry with specific antibodies.
• Proto-Pyrenoid Visualization: Analyze chloroplasts from transgenic lines using confocal microscopy. The dense, phase-separated proto-pyrenoid structures can be identified by the co-localization of fluorescently tagged Rubisco and EPYC1.

Validation: Successful implementation results in the formation of a single, dense phase-separated droplet of Rubisco and EPYC1 within the chloroplast, a structure absent in wild-type plants [74].

Protocol: Enhancing Heavy Metal Phytoextraction with Engineered Microalgae

This protocol outlines a synthetic biology approach to improve microalgae's capacity for heavy metal uptake [72].

Objective: To genetically engineer Chlamydomonas reinhardtii for increased tolerance and accumulation of heavy metals like mercury (Hg) or chromium (Cr).

Key Reagents:

• Algal Strain: Chlamydomonas reinhardtii CC-503 cw92 mt+.
• Engineering Tool: CRISPR/Cas9 system tailored for C. reinhardtii or modular cloning (MoClo) toolkit parts.
• Genetic Parts:
  • Promoter: A strong, constitutive promoter (e.g., HSP70A/RBCS2).
  • Gene of Interest: Sequences encoding metallothioneins (metal-binding proteins) or metal transporters (e.g., ZRT/IRT-like Protein (ZIP) family).
  • Selection Marker: An antibiotic resistance gene (e.g., Paromomycin resistance AphVII).
• Growth Medium: Tris-Acetate-Phosphate (TAP) medium, with and without added heavy metal salts (e.g., HgCl₂, K₂CrO₄).

Methodology:

• Construct Assembly: Assemble the expression cassette using a modular cloning system like the C. reinhardtii MoClo toolkit. The cassette should contain the promoter, gene of interest, and selection marker.
• Algal Transformation: Deliver the linearized DNA construct into C. reinhardtii cells via glass bead agitation or electroporation.
• Mutant Selection: Plate transformed cells on TAP agar plates containing the relevant antibiotic. Isolate single colonies after 7-10 days.
• Screening and Characterization:
  • Molecular Confirmation: Validate gene integration and expression in transgenic lines using PCR, qRT-PCR, and Western blot.
  • Tolerance Assay: Grow wild-type and transgenic lines in TAP liquid medium spiked with sub-lethal and lethal concentrations of the target heavy metal. Monitor growth kinetics by measuring optical density (OD₆₈₀) over 5-7 days.
  • Uptake Quantification: Harvest cells from metal-containing media, wash, and digest with nitric acid. Measure intracellular metal concentration using Inductively Coupled Plasma Mass Spectrometry (ICP-MS).

Validation: Successful engineering is indicated by transgenic algae exhibiting significantly higher growth rates and final biomass in metal-spiked media, coupled with a higher intracellular concentration of the target metal compared to wild-type controls [72].

Signaling Pathways and Metabolic Engineering Workflows

The following diagrams, generated using DOT language, illustrate the core metabolic and genetic engineering strategies discussed.

Metabolic Strategies to Enhance Photosynthetic Carbon Fixation

Diagram 1: Metabolic engineering strategies to overcome RuBisCO limitation. Pathways contrast the native C3 plant cycle with two engineering approaches: introducing an algal Carbon Concentrating Mechanism (CCM) and creating a synthetic RuBisCO bypass using PEPC and the MOG cycle [73] [75] [74].

Genetic Engineering Workflow for Pollutant-Uptake in Microalgae

Diagram 2: A generalized genetic engineering workflow for enhancing pollutant uptake or degradation in microalgae. The process involves identifying key genes, assembling genetic constructs, transforming algae, and rigorously screening for improved performance [72].

The Scientist's Toolkit: Key Research Reagents

This section details essential reagents, components, and organisms used in the experiments cited, providing a resource for researchers aiming to work in this field.

Table 3: Key Research Reagents and Organisms

Reagent / Organism	Type/Function	Application Example
Chlamydomonas reinhardtii	Model green alga	Source of CCM genes (EPYC1, LCIA); chassis for bioremediation engineering [75] [72] [74]
Arabidopsis thaliana	Model higher plant (C3)	Chassis for introducing algal CCM components and testing pyrenoid formation [76] [74]
EPYC1 (Linker Protein)	Algal protein with multiple Rubisco-binding sites	"Molecular glue" for inducing phase separation of Rubisco into pyrenoid-like structures in plants [74]
RuBisCO (Hybrid Engineered)	Key carbon-fixing enzyme	Engineered with algal sequences to bind EPYC1, enabling condensate formation in plant chloroplasts [74]
Phosphoenolpyruvate Carboxylase (PEPC)	High-affinity CO₂ fixing enzyme	Proposed replacement for RuBisCO in synthetic carbon fixation cycles (e.g., MOG cycle) [73]
Carbonic Anhydrase (CA)	Enzyme interconverting CO₂ and HCO₃⁻	Enhancing mesophyll conductance in plants; critical component in algal CCMs [75] [76]
CRISPR/Cas9 Systems	Gene editing tool	Precise genome modification in both microalgae and higher plants for knockout/knock-in of target genes [72]
Modular Cloning (MoClo) Toolkits	Standardized genetic parts assembly	Rapid and efficient construction of complex genetic circuits in microalgae (e.g., C. reinhardtii toolkit) [72]
Metallothioneins	Cysteine-rich metal-binding proteins	Overexpression in microalgae to enhance sequestration and tolerance of heavy metals [72]

The pursuit of advanced air revitalization systems for closed environments and extraterrestrial habitats has intensified the search for highly efficient biological solutions. Within this context, a critical comparison between microalgae and traditional higher plants is essential, focusing on the core engineering challenges of hydrodynamics, contamination control, and system maintenance. Microalgae, with their high photosynthetic efficiency and rapid growth rates, present a promising alternative to higher plants. However, their deployment in practical systems is governed by the complex interplay of fluid dynamics, susceptibility to biological contamination, and operational upkeep requirements. This guide objectively compares the performance of microalgae-based systems against higher plants, drawing on current experimental data to inform researchers and scientists in the field of bioregenerative life support systems.

Performance Comparison: Microalgae vs. Higher Plants

The design and operation of biological air revitalization systems require a fundamental understanding of the performance characteristics of the primary photosynthetic organisms. The table below provides a structured, data-driven comparison between microalgae and higher plants, focusing on key parameters relevant to system efficiency, scalability, and operational demands.

Table 1: Quantitative performance comparison of microalgae and higher plants for air revitalization applications.

Performance Parameter	Microalgae Systems	Higher Plant Systems	Comparison Implications
Carbon Sequestration Efficiency	1.3 - 1.83 kg CO₂ per kg biomass produced [7] [77]; 10-50 times greater than terrestrial plants [7]	Varies by species; generally lower per unit area and time	Microalgae offer superior CO₂ fixation potential in space-constrained environments.
Oxygen Production	Contributes ~50% of global O₂ production [15]	Significant, but lower aerial productivity	Microalgae are highly efficient O₂ producers on an areal basis.
Hydrodynamic Dependence	Extremely high; mixing is critical for light/dark cycles, nutrient distribution, and gas exchange [78] [79]	Primarily driven by transpiration and root hydration; less intense mixing required	Microalgae systems demand more complex and energy-intensive hydrodynamic design.
Contamination Risk & Type	High risk of culture crash from invasive microalgae, bacteria, and predators [78] [80]	Risk from pests, fungi, and pathogens; generally more manageable	Microalgae contamination is often irreversible, requiring full system sterilization.
Nutrient Source & Maintenance	Can utilize wastewater and flue gas; harvesting required every few days [7] [80]	Typically require purified water and soil/fertilizer; harvest cycles are weeks to months	Microalgae enable resource recovery but need frequent, continuous harvesting.
Growth Rate & Space Requirement	Very high growth rate; high productivity in compact photobioreactors [24] [15]	Slower growth; requires larger soil volume and canopy space for equivalent yield	Microalgae are superior for applications with severe mass and volume constraints.
Water Usage	Can be cultivated in brackish or wastewater; minimal freshwater consumption [15]	Generally require high-quality freshwater for irrigation	Microalgae systems align better with closed-loop water recycling paradigms.

Hydrodynamic Challenges in Microalgae Cultivation

Hydrodynamics is a pivotal factor influencing the scalability and productivity of microalgae cultivation systems, directly impacting mass transfer, light exposure, and shear stress.

Experimental Insights and Protocols

Research on Synechococcus HS-9 cultivation in a Rectangular Airlift Photobioreactor with Baffles (RAPBR-Bs) quantified the relationship between hydrodynamics and growth. The experimental protocol involved [79]:

• Bubble Dynamics Analysis: High-speed photography and video were used to measure bubble properties, including velocity (0.0064 m/s) and diameter (720 μm).
• Hydrodynamic Parameter Calculation: Key non-dimensional numbers were computed, such as Reynolds (Re = 4.51), Eötvös (Eo = 0.0126), and Weber (We = 6.85 × 10⁻⁵) numbers.
• Mass Transfer Measurement: The volumetric mass transfer coefficients for oxygen (kLa O₂ = 0.114 s⁻¹) and carbon dioxide (kLa CO₂ = 0.099 s⁻¹) were determined.
• Biomass Monitoring: Optical density and dry biomass weight were tracked over time, revealing optimal growth at day 13 with a biomass yield of 3.226 g under optimal mixing and aeration.

In raceway pond systems, hydrodynamic optimization focuses on overcoming challenges like dead zones and ensuring adequate flow velocity. Studies employ Computational Fluid Dynamics (CFD) to model flow fields and optimize paddle wheel design and pond geometry to minimize energy consumption while achieving sufficient mixing to prevent sedimentation and ensure cyclic light exposure [78].

Comparative Hydrodynamic Workflow

The fundamental difference in hydrodynamic requirements between microalgae and plant systems is illustrated below.

Contamination Control and System Maintenance

Contamination and maintenance are critical determinants of operational longevity and reliability.

Contamination Dynamics and Protocols

Microalgae cultures in open systems are highly susceptible to invasion by unwanted microalgae species, bacteria, and predators like rotifers, which can lead to complete culture collapse within days. A meta-analysis of pilot-scale High-Rate Algal Ponds (HRAPs) confirmed that operating with real wastewater leads to dynamic microalgae-bacteria consortia, where the community composition constantly shifts in response to environmental fluctuations [80]. Control strategies include:

• Prevention: Using closed photobioreactors instead of open ponds drastically reduces contamination risk [79].
• Culture Management: Maintaining a high inoculation density and a low hydraulic retention time (HRT) can help the desired species outcompete invaders.
• Chemical Treatment: The use of pesticides or antibiotics is generally avoided as it compromises the biomass for subsequent applications.

In contrast, higher plant systems face threats from insects, fungi, and soil-borne diseases. Control methods include integrated pest management, sterile growth chambers, and fungicides. The "maintenance" in plant systems often involves pruning, trellising, and pollination, which are labor-intensive but mechanically simpler than microalgae harvesting.

Maintenance Requirements and Experimental Data

Maintenance in microalgae systems is characterized by frequent, automated processes. A key finding from pilot-scale research is that biomass growth rates at scale (~0.54 day⁻¹) are roughly half those observed in controlled laboratory experiments, highlighting a significant scaling bottleneck often related to suboptimal maintenance and hydrodynamic conditions [80].

Table 2: Comparison of operational and maintenance requirements.

Maintenance Activity	Microalgae Systems	Higher Plant Systems
Harvesting / Cropping	Continuous (every 2-5 days); requires centrifugation/flocculation [80]	Cyclical (weeks/months); manual or mechanical harvesting
Nutrient Supply	Continuous dosing; can be automated with sensors	Batch fertilization or continuous hydroponic solution replenishment
System Cleaning	Frequent biofouling control in reactors and pipes; sterilization between batches	Periodic cleaning of hydroponic systems; soil replacement if used
Water Replenishment	Low due to minimal evaporation from closed systems	High due to plant transpiration
Process Monitoring	Requires daily checks of OD, pH, and contamination via microscopy	Monitoring for pest/disease, nutrient deficiencies, and water status

The Scientist's Toolkit: Essential Research Reagents and Materials

The experimental study of these systems relies on a suite of specialized reagents and equipment.

Table 3: Key research reagent solutions and materials for microalgae and plant air revitalization research.

Reagent / Material	Function in Research	Application Example
BG-11 / BBM Media	Standardized nutrient medium providing essential macronutrients (N, P) and micronutrients for axenic microalgae culture.	Serves as a controlled, synthetic growth medium to establish baseline growth kinetics without the variability of wastewater [79].
Tracer Dyes (e.g., Rhodamine)	Used to characterize hydrodynamic performance and determine parameters like hydraulic retention time (HRT) distribution and dead zones.	Injected into raceway ponds or constructed wetlands to visualize flow paths and measure hydraulic efficiency (λ) [81] [82].
Computational Fluid Dynamics (CFD) Software	Numerical modeling tool for simulating fluid flow, mass transfer, and shear stress within photobioreactors or root zones.	Used to optimize raceway pond baffle design, sparger placement in airlift reactors, and predict mixing times [81] [79].
Fluorescent Probes for kLa Measurement	Enables experimental determination of the volumetric mass transfer coefficient (kLa), a critical parameter for gas (O₂, CO₂) exchange efficiency.	Quantifying how reactor design (e.g., with/without baffles) improves the mass transfer of CO₂, which is often the growth rate-limiting factor [79].
CO₂ Gas Calibration Standards	Pre-mixed gases of known CO₂ concentration for calibrating sensors and supplying carbon for phototrophic growth experiments.	Essential for maintaining optimal CO₂ levels in photobioreactors and for studies on CO₂ bio-fixation rates from flue gas simulants [77].
DNA Extraction Kits & PCR Primers	For molecular identification and monitoring of microbial community composition (e.g., algae, bacteria, contaminants) in culture.	Tracking population dynamics in open pond systems and identifying specific contaminant species during a culture crash [80].

Carbon Assimilation Pathways

The core biochemical process driving air revitalization is carbon fixation. The pathway divergence between microalgae and higher plants is a fundamental differentiator.

The choice between microalgae and higher plants for air revitalization involves a direct trade-off between performance and engineering complexity. Microalgae systems demonstrate superior metrics in carbon sequestration efficiency, oxygen production rate, and resource utilization (water, nutrients from waste streams), making them theoretically ideal for mass- and volume-critical applications. However, this high performance is contingent upon successfully overcoming significant system design challenges related to energy-intensive hydrodynamics, high susceptibility to contamination, and frequent, complex maintenance protocols. Higher plant systems, while less efficient on an areal basis, offer lower hydrodynamic complexity and potentially more robust, manageable growth cycles. The optimal path forward may not be a choice of one over the other, but rather the development of integrated, hybrid life support systems that leverage the unique strengths of both biological agents. Future research should prioritize the development of more contamination-resistant microalgae strains, energy-efficient hydrodynamic designs, and automated maintenance protocols to unlock the full potential of microalgae-based air revitalization.

Data-Driven Comparison: Quantifying Air Revitalization Performance

The escalating concentration of atmospheric carbon dioxide (CO2) is a primary driver of global climate change, necessitating the development of efficient biological carbon sequestration strategies. Within this context, the competition between microalgae and terrestrial plants for the most effective air revitalization system is a critical area of research. This guide provides an objective, data-driven comparison of the CO2 fixation performance of these two biological systems. It is designed to support researchers and scientists in evaluating each system's potential by synthesizing current performance benchmarks, detailing standard experimental methodologies, and outlining the essential tools for investigation. The comparative analysis focuses on fixation rates, photosynthetic efficiency, and the synergistic potential of each system within a biorefinery model, providing a scientific foundation for future research and development in carbon capture technologies.

Performance Benchmarking: Quantitative Data Comparison

Direct quantitative comparison reveals distinct performance advantages and trade-offs between microalgae and terrestrial plant systems for CO2 fixation. The data, synthesized from recent studies, are summarized in the table below for clear benchmarking.

Table 1: Performance Benchmarking: Microalgae vs. Terrestrial Plants

Performance Metric	Microalgae	Terrestrial Plants	Notes & Context
CO2 Fixation Rate	1.0–3.7 g CO₂/L/day [26]; ~2.4 g CO₂/L/day (theoretical based on biomass) [83]	~157 Pg C/year (Global GPP) [84]	Microalgae rates are for optimized bioreactors; terrestrial plant rate is a refreshed global estimate.
Biomass Productivity	Up to 280 tons dry biomass/ha/year [85]	~27.8 Mg/ha/year (Hybrid Poplar, above-ground) [86]; ~10-11 Mg/ha/year (Maize, above-ground) [86]	1 Mg = 1 metric ton. Microalgae productivity is theoretical under ideal conditions.
Photosynthetic Efficiency	10 to 50 times higher than terrestrial plants [85] [7]	Used as baseline (1x)	Attributed to microalgae's Carbon Concentrating Mechanism (CCM) and simpler cellular structure [7].
Carbon Sequestration Efficiency (CDSE)	Up to 30.0% (Tribonema minus at 1.5% CO2) [83]	Not directly comparable; long-term sequestration depends on biomass use (e.g., wood products).	CDSE is a specific metric for bio-capture in controlled systems.
Land Use	Can use non-arable land and wastewater, avoiding competition with crops [85] [7]	Requires arable land, leading to potential competition with food production.	A major strategic advantage for microalgae.
Key Advantages	High growth rate, continuous harvest, produces high-value co-products (biofuels, pigments) [26] [85]	Simpler large-scale cultivation, massive existing infrastructure, long-term carbon storage in woody biomass [86]

The data indicates that microalgae possess a superior carbon fixation rate per unit volume or area and significantly higher photosynthetic efficiency due to their carbon-concentrating mechanisms and ability to utilize a greater fraction of available light [85] [7]. Furthermore, their cultivation does not compete with food production for arable land. Conversely, terrestrial plants, particularly woody perennials, play an irreplaceable role in global carbon cycles and offer the benefit of long-term carbon sequestration in woody biomass and root systems, which can store carbon for decades to centuries [86]. The recent upward revision of global terrestrial GPP to 157 petagrams of carbon per year underscores the immense, albeit diffuse, capacity of natural ecosystems [84].

Experimental Protocols for Measuring CO2 Fixation

Accurately benchmarking CO2 fixation requires robust and standardized experimental protocols. The methodologies differ significantly between microalgae and plants, reflecting their distinct biological and cultivation contexts.

Microalgae Cultivation and Carbon Fixation Analysis

Microalgal CO2 fixation is typically measured under controlled laboratory conditions in photobioreactors (PBRs). The following protocol outlines a standard method for evaluating carbon dioxide sequestration efficiency (CDSE).

• 1. Strain Selection and Pre-culture: Select and inoculate target strains (e.g., Desmodesmus armatus, Tribonema minus) into a nutrient-rich medium like Bold's Basal Medium (BBM 3N). Pre-culture for several days under optimal conditions (e.g., 22.5°C, 150 μmol photons/m²/s) on an orbital shaker to obtain an exponentially growing inoculum [83].
• 2. Controlled Cultivation in Photobioreactors: Transfer the inoculum to a Laboratory System for Intensive Cultivation (LSIC) or column PBRs. Key parameters must be strictly controlled:
  • Temperature: Maintain at 27.0 ± 0.6°C [83].
  • Illumination: Provide continuous 24-hour illumination at 300 μmol photons/m²/s using white LED lights [83].
  • CO2 Supplementation: Continuously bubble the culture with air (0.04% CO2) or a CO2-enriched air mixture (e.g., 1.5% CO2) at a fixed flow rate (e.g., ~0.2 L/min) [83]. Advanced PBRs may employ optimized baffles to enhance gas-liquid mixing and increase bubble retention time, thereby boosting CO2 utilization [87].
• 3. Biomass and Carbon Monitoring: Sample the culture periodically (e.g., days 0, 3, 6, 9). Key measurements include:
  • Optical Density (OD750): For rapid assessment of biomass growth [83].
  • Dry Biomass Concentration: Filter a known culture volume, dry the biomass, and weigh it [83].
  • Organic Carbon Concentration: Analyze using elemental analysis or chemical oxygen demand (COD) methods [83].
• 4. Data Calculation: The Carbon Dioxide Sequestration Efficiency (CDSE) is calculated based on the carbon content in the biomass relative to the total carbon input from the CO2-enriched air supply [83].

Terrestrial Plant Photosynthesis Measurement

For terrestrial plants, the gross primary production (GPP) representing total CO2 fixed via photosynthesis is measured at scales from the leaf to the globe.

• 1. Leaf-Level Gas Exchange: Using portable infrared gas analyzer (IRGA) systems, measure the net CO2 flux into a single leaf under controlled light, CO2, and humidity conditions. This provides direct data on photosynthetic rates but requires upscaling [84].
• 2. Ecosystem-Level Eddy Covariance: This micrometeorological technique uses towers to measure the vertical flux of CO2 between the ecosystem and the atmosphere. It provides integrated, continuous data on net ecosystem exchange (NEE), which is used to model GPP over large areas [84].
• 3. Global GPP Estimation via Carbonyl Sulfide (OCS) Proxy: A novel method uses atmospheric carbonyl sulfide (OCS) as a proxy. OCS follows a similar pathway to CO2 during plant diffusion but is not respired. By modeling the global atmospheric budget and uptake of OCS, scientists can infer GPP more accurately, leading to revised estimates like the 157 petagrams of carbon per year [84].

The following workflow diagram illustrates the logical relationship and progression of these key methodologies from controlled laboratory studies to global-scale modeling.

Diagram 1: Experimental Workflow for CO2 Fixation Benchmarking. This diagram outlines the parallel methodological pathways for quantifying CO2 fixation in microalgae (controlled bioreactors) and terrestrial plants (field to global scales), culminating in a comparative analysis.

Research Reagent Solutions and Essential Materials

Conducting research in this field requires specific reagents, biological materials, and specialized equipment. The table below details key solutions essential for experimental work in microalgae and terrestrial plant CO2 fixation studies.

Table 2: Essential Research Reagents and Materials

Category	Item	Function/Application	Representative Example / Composition
Culture Media	Bold's Basal Medium (BBM 3N)	A standardized nutrient solution for cultivating a wide range of microalgae in controlled experiments.	Modified BBM with 3x nitrogen (NaNO₃ - 750 mg/L) and vitamin B12 supplementation [83].
Biological Materials	Novel Microalgae Strains	Bioprospecting for strains with high CO2 tolerance and fixation efficiency is crucial for technology development.	Desmodesmus armatus ARC-06 (low CO2 tolerant), Tribonema minus ARC-10 (high CO2 performer) [83].
Gas Systems	CO2 Enriched Air Supply	Provides the primary inorganic carbon source for microalgae cultivation in PBRs, simulating flue gas or high-CO2 environments.	Pre-mixed air with 1.5% (v/v) CO2, delivered via bubbling or specialized spargers [83].
Photobioreactors	Column PBR with Baffles	The core cultivation vessel. Internal structures like Portable Conical Helix Baffles (PCHB) enhance mixing and CO2 mass transfer.	3D-printed round PCHB to generate spiral flow vortices, increasing biomass yield by up to 33% [87].
Analytical Tools	Carbonyl Sulfide (OCS)	A atmospheric tracer used as a proxy to measure gross primary production (GPP) in terrestrial ecosystems at a global scale.	Tracking OCS uptake by plants, which follows a similar diffusion path as CO2 but is not respired [84].
Analytical Instruments	LI-COR Quantum Radiometer	Precisely measures photosynthetically active radiation (PAR), a critical parameter for standardizing light conditions in PBRs and field studies.	Li-189 radiometer with LI-190SA sensor for measuring μmol photons/m²/s [83].

This comparison guide elucidates that the choice between microalgae and terrestrial plants for CO2 fixation is not a simple matter of superiority but one of application context. Microalgae demonstrate unparalleled rate-based efficiency and biorefinery potential within controlled, high-intensity systems, making them ideal for targeted carbon capture from industrial point sources and simultaneous production of high-value biofuels and chemicals. Their performance is heavily dependent on sophisticated engineering and optimization of cultivation systems [26] [87] [88]. In contrast, terrestrial plants, with their vast spatial scale and existing biomass, represent a fundamental, nature-based solution for global atmospheric carbon management, with recent data confirming their critical role is even larger than previously estimated [84]. The future of biological air revitalization likely lies not in selecting one system over the other, but in strategically deploying both to create synergistic, multi-scale carbon sequestration networks that address the climate crisis from laboratory to landscape.

The escalating challenge of indoor and environmental air pollution has intensified the search for effective and sustainable bioremediation strategies. Among the most promising avenues is the use of photosynthetic organisms, primarily drawing a comparison between traditional higher plants and emerging microalgae technologies. The concept of using higher plants for air purification gained popular traction following seminal NASA studies, but recent scientific reviews have questioned their efficacy under real-world conditions. Concurrently, advances in biotechnology have highlighted the potential of microalgae—diverse, fast-growing photosynthetic microorganisms—in targeted pollutant removal. This guide provides a objective, data-driven comparison of these two biological systems, focusing on their efficacy, mechanisms, and practical applications for removing formaldehyde, volatile organic compounds (VOCs), and particulate matter (PM). The analysis is framed within the broader research context of air revitalization efficiency, providing scientists and researchers with a critical evaluation of experimental data and methodologies.

The following tables synthesize quantitative data on the pollutant removal capabilities of higher plants and microalgae, drawing from controlled experiments and performance analyses.

Table 1: Efficacy in Removing Formaldehyde and VOCs

System / Organism	Target Pollutant	Experimental Concentration	Removal Efficiency / Rate	Key Experimental Conditions
Dynamic Botanical Air Filter (DBAF) [89]	Formaldehyde	< 50 ppb	~60% single-pass efficiency	Dynamic air flow through root bed; sustained over 10 months.
Static Potted Plant [89]	Formaldehyde	10 ppm	Negligible removal	Sealed chamber; no active air flow through growth media.
Active Green Wall System [89]	Formaldehyde	Not Specified	Clean Air Delivery Rate (CADR): 232–759 m³/h/m²	Dynamic botanical air filtration system.
Microalgae (General) [26]	CO₂ (as a VOC metric)	N/A	Carbon fixation: 1.0–3.7 g CO₂/L/day	Optimized photobioreactor conditions.

Table 2: Efficacy in Removing Other Pollutants

System / Organism	Target Pollutant	Experimental Context	Removal Mechanism & Notes
Microalgae [19]	Spent Oil Waste (SOW), Hydrocarbons	Biodegradation of industrial waste	Adsorption, bioaccumulation, and biotransformation into CO₂ and water.
Microalgae [90]	Heavy Metals	Wastewater treatment (Phycoremediation)	Extracellular and intracellular mechanisms; potential for integrated systems.
Houseplants [91]	Particulate Matter (PM)	Real-world indoor environments	Leaf surface accumulation; effectiveness limited without high plant density.

Detailed Experimental Protocols and Mechanisms

Removal of Formaldehyde and VOCs

a) Dynamic Botanical Air Filtration (DBAF) for Formaldehyde Removal

• Objective: To investigate the fundamental mechanisms of formaldehyde removal in an active botanical filtration system and differentiate the roles of plant leaves, the wet sorbent bed, and root-zone microbes [89].
• Methodology:
  • Group A (Static Potted Plant): A potted plant was placed in a sealed chamber with an initial formaldehyde concentration of 10 ppm. No air was passed through the root bed, isolating the effect of the plant's phyllosphere [89].
  • Group B (Microbial Community): An experimental setup with air flow passing through a medium containing only the microbial community from the root bed, excluding the plant itself [89].
  • Group C (Full DBAF System): A dynamic botanical air filter with air actively drawn through the plant's root bed [89].
  • Analysis: The decay rate of formaldehyde concentration in each chamber was measured over time and compared to an inert tracer gas (SF₆) to account for air leakage [89].
• Key Findings: The study concluded that dynamic air flow through the root bed is essential for meaningful formaldehyde removal. The root zone microbial community was identified as the primary agent of degradation, with moisture playing a critical role in maintaining microbial activity. The leaves of the static potted plant contributed negligibly to pollutant removal [89].

b) Limitations of Higher Plants in Real-World Settings

A critical meta-analysis reviewed decades of research on potted plants and VOC removal. It concluded that to replicate the purification effects observed in small-scale lab studies in a typical office or home environment, one would require an impractical density of 10 to 1,000 plants per square meter of floor space. In a standard 1,500 square foot home, this translates to approximately 680 plants. The analysis further noted that in real buildings, natural or mechanical ventilation dominates VOC removal, with plants providing a negligible additive effect [91].

c) Microalgae Mechanisms for Hydrocarbon Degradation

Microalgae degrade complex hydrocarbons like those in spent oil waste through a multi-step mechanism:

• Adsorption: Hydrocarbons adhere to the surface of the microalgal cell [19].
• Bioaccumulation & Emulsification: The hydrocarbons are taken up into the cell via active or passive transport [19].
• Biotransformation: Within the cell, enzymatic processes break down the complex hydrocarbons into harmless compounds such as carbon dioxide and water [19].

This process, driven by fast-growing microalgae with high photosynthetic efficiency, can be optimized in controlled photobioreactors for waste treatment and carbon sequestration [26] [19].

Removal of Particulate Matter and Heavy Metals

a) Higher Plants and Particulate Matter (PM)

• Mechanism: The primary pathway for PM removal by higher plants is the dry deposition of particles onto the leaf surface. The complex surface topography of leaves can trap and retain PM [91].
• Limitations: The effectiveness of this process in improving indoor air quality is limited. Dust accumulated on leaves can be re-suspended into the air by disturbances. Furthermore, overwatered potted plants can promote mold growth in the soil, which itself becomes a source of airborne biological pollutants [91].

b) Microalgae and Heavy Metal Phycoremediation

• Mechanism: Microalgae remove heavy metals through extracellular and intracellular processes, including biosorption onto the cell wall and bioaccumulation within the cell. They exhibit various self-defense mechanisms to resist metal toxicity [90].
• Applications: This capability is primarily harnessed for wastewater treatment. Research is focused on developing integrated systems to make phycoremediation a more economical and sustainable technology for metal recovery and water purification [90].

Signaling Pathways and Workflows

The experimental approaches and fundamental mechanisms can be visualized in the following workflows.

The Scientist's Toolkit: Research Reagent Solutions

Table 3: Essential Reagents and Materials for Air Bioremediation Research

Reagent / Material	Function in Research	Example Application
Bold's Basal Medium (BBM)	Standardized nutrient medium for culturing freshwater microalgae and cyanobacteria.	Cultivation of Chlorella vulgaris and Arthrospira platensis for pollutant tolerance tests [92].
Gas Chromatography-Mass Spectrometry (GC-MS)	High-sensitivity identification and quantification of volatile organic compounds (VOCs).	Profiling volatiles in microalgae biomass [92] or measuring VOC removal in chamber studies [89].
Scanning Electron Microscopy with Energy Dispersive X-ray Spectroscopy (SEM-EDX)	Visualization of surface morphology and elemental composition of samples.	Characterizing interaction between microplastics and microalgae [21] or analyzing biofilm formation on filters.
Fourier Transform Infrared Spectroscopy (FT-IR)	Identification of molecular bonds and functional groups in a sample.	Polymer characterization of microplastics and studying their chemical interaction with biological systems [21].
Ceramic Membrane Filters	High-stability filtration for harvesting microalgae biomass from growth media.	Dewatering mixed microalgae cultures grown in wastewater for subsequent analysis or processing [93].
Solid Phase Extraction (SPE)	Fractionation and purification of complex mixtures of organic compounds from samples.	Separating volatile compounds from microalgal biomass into fractions for improved GC-MS identification [92].

The comparative analysis reveals a clear functional distinction between higher plants and microalgae in air pollutant remediation. Higher plants, particularly in static potted configurations, demonstrate negligible removal of gaseous pollutants like formaldehyde in real-world settings, with their efficacy becoming significant only in densely packed, active systems that leverage the root-zone microbiome. Their primary mechanical action for particulate matter is limited to leaf surface deposition. In contrast, microalgae operate as versatile cellular biofactories, capable of actively degrading a wide spectrum of pollutants—from VOCs and complex hydrocarbons to heavy metals—through intracellular and extracellular processes. Their high photosynthetic efficiency and adaptability make them exceptionally suited for engineered systems like photobioreactors for wastewater treatment and targeted air revitalization. While higher plants offer psychological and aesthetic benefits, microalgae present a technologically superior pathway for efficient, scalable, and sustainable bioremediation, meriting continued research and development for environmental applications.

Analysis of Oxygen Production and Impact on Indoor Relative Humidity

The pursuit of effective air revitalization systems for confined environments has intensified research into biological approaches for oxygen production and carbon dioxide removal. Among these, microalgae-based systems and higher plants represent two fundamentally different biological strategies. Microalgae, comprising diverse groups of photosynthetic microorganisms including eukaryotic green algae and prokaryotic cyanobacteria, offer distinctive advantages for engineered air revitalization systems [94]. These organisms operate with significantly higher photosynthetic efficiency than terrestrial plants due to their simplified cellular structure and capacity for full biomass utilization [3]. With indoor air quality emerging as a critical health concern—particularly given that Americans spend approximately 90% of their time indoors—the need for effective biological air purification has never been greater [95]. This analysis provides a comparative assessment of microalgae versus higher plants for air revitalization, focusing specifically on oxygen production capabilities and associated impacts on indoor relative humidity, with supporting experimental data for researchers and scientific professionals.

Quantitative Comparison: Oxygen Production and Environmental Impact

Table 1: Comparative Performance Metrics for Air Revitalization Systems

Parameter	Microalgae Systems	Higher Plants	Measurement Context
Oxygen Production Rate	5-10× higher than terrestrial plants [94]	Baseline	Per unit biomass
CO2 Sequestration Efficiency	10-50× higher than terrestrial plants [85] [7]	Baseline	Per unit area
Biomass Productivity	80-280 tons/ha/year [85] [3]	Significantly lower	Annual yield
Water Consumption	Reduced (closed systems minimize evaporation) [94]	Higher due to evaporation and infiltration	Per unit biomass
Humidity Contribution	Actively managed via closed systems	Passive transpiration	Impact on indoor environment
Space Requirements	Compact photobioreactors (vertical integration possible)	Significant horizontal space required	Footprint per oxygen unit
Light Utilization Efficiency	3-9% of theoretical maximum [3]	Typically lower	Photosynthetic efficiency

Table 2: Microalgae Species Performance Characteristics

Species	Specific Applications	Oxygen Production Characteristics	Growth Requirements
Chlorella vulgaris	Air purification, nutrition	High biomass productivity (up to 6.48×10^7 cells/mL) [96]	Controlled photoperiod (16:8 light:dark optimal) [96]
Spirulina (Arthrospira)	Nutritional supplements, air revitalization	Efficient photosynthesis under high pH	Tolerates high pH conditions [94]
Haematococcus pluvialis	Astaxanthin production	Standard photosynthetic rate	Performs well in closed reactors [94]
Chlamydomonas reinhardtii	Model organism for research	Well-characterized photosynthetic mechanism	Adaptable to various photobioreactors [94] [3]
Dunaliella salina	β-carotene production	Efficient under high salinity	Tolerates high salt concentrations [94]

Experimental Protocols for Performance Evaluation

Photobioreactor Cultivation Methodology

The standardized protocol for assessing microalgae performance employs controlled photobioreactor systems. Experiments with Chlorella vulgaris demonstrate the critical importance of precise light management in optimizing oxygen production [96]. The methodology involves cultivation in 3.5L Phyto tank systems with integrated LED lighting, controlled airflow, and sterile conditions maintained at 22.0±2.0°C [96]. Biomass concentration is quantified using automated fluorescence cell counters and spectrophotometry (measuring optical density at 682nm, the most sensitive wavelength for C. vulgaris) [96]. The 16:8-hour light-dark photoperiod has been experimentally verified to yield the highest biomass concentration (6.48×10^7±0.50 cells/mL with OD 1.165), significantly outperforming continuous illumination or other photoperiods [96]. This protocol provides a standardized approach for comparing oxygen production capabilities across different microalgae strains.

Humidity Impact Assessment Protocol

The interaction between photosynthetic organisms and indoor humidity requires controlled experimental assessment. Research indicates that relative humidity directly influences microbial growth on surfaces, with studies showing no significant algae growth below 98% relative humidity from an engineering standpoint [97]. The experimental protocol involves accelerated growth tests under precisely controlled humidity conditions using environmental chambers. Temperature and humidity sensors are deployed to monitor conditions, with algal growth quantified through image analysis of covered area and biomass measurements [97]. These experiments have successfully modeled biofouling using a modified Avrami's law, providing a mathematical framework for predicting growth under various humidity conditions [97]. This methodology allows researchers to differentiate between the passive humidity contribution of plants and the actively managed humidity in closed photobioreactor systems.

Microalgae Photosynthesis and Humidity Relationship Diagram: This diagram illustrates the interconnected processes of light capture, photosynthetic reactions, and humidity relationships in microalgae-based air revitalization systems.

The Scientist's Toolkit: Essential Research Reagents and Materials

Table 3: Essential Research Materials for Microalgae Air Revitalization Studies

Reagent/Equipment	Function/Application	Specific Examples
Photobioreactors	Controlled cultivation environment	Flat panel airlift reactors, tubular reactors, Phyto tank systems (3.5L) [94] [96]
Lighting Systems	Photosynthesis driver	LED arrays (warm-white fluorescent, 3000 Lux) [96], specific wavelengths [59]
Culture Media	Nutrient source for growth	Bold Basal Medium [96], N-8 medium [98]
Analytical Instruments	Growth and oxygen measurement	Spectrophotometers (682nm for C. vulgaris) [96], automated cell counters [96]
Environmental Monitors	Humidity and temperature tracking	Relative humidity sensors, temperature loggers [97]
Gas Exchange Analyzers	Oxygen/CO2 measurement	Systems for quantifying photosynthetic rates [11]
Algal Strains	Experimental subjects	Chlorella vulgaris UTEX 2714 [96], Spirulina, Chlamydomonas reinhardtii [94]

Comparative Analysis and Research Implications

Oxygen Production Efficiency

Microalgae demonstrate substantially higher oxygen production capabilities compared to higher plants on a per-biomass basis. This efficiency stems from their simplified cellular structure and dedication of nearly all biomass to photosynthetic activity, unlike higher plants that allocate significant resources to structural components like stems and roots [3]. The theoretical maximum photosynthetic efficiency for microalgae ranges between 9-10%, corresponding to approximately 80g of biomass/m²/day or 280 tons/ha/year [3]. In practical industrial-scale photobioreactors, microalgae typically achieve 3-5% light conversion efficiency, still significantly exceeding the performance of most terrestrial plants [3]. This enhanced efficiency makes microalgae particularly valuable for space-constrained indoor environments where maximizing oxygen production per unit area is critical.

Humidity Management Considerations

The impact on indoor relative humidity represents a crucial differentiation between microalgae and plant-based systems. Closed photobioreactors used for microalgae cultivation effectively isolate humidity generation from the indoor environment, allowing for active management of moisture levels [94]. In contrast, higher plants continuously release water vapor through transpiration processes, directly increasing indoor humidity levels [97]. Research has demonstrated that algal growth requires high humidity environments (greater than 98% relative humidity) [97], but closed-system cultivation prevents this humidity from affecting indoor spaces. This controlled humidity profile presents significant advantages for maintaining comfortable indoor environments while achieving high oxygen production rates.

Applications in Specialized Environments

The superior oxygen production and controllable humidity impact of microalgae systems make them particularly suitable for specialized environments where precise atmospheric control is essential. Space mission applications represent a prime example, where microalgae are integrated into Bioregenerative Life Support Systems (BLSS) for simultaneous air revitalization, water recycling, and food production [11]. Current Environmental Control and Life Support Systems (ECLSS) on the International Space Station utilize physicochemical processes that achieve only partial resource recovery, with microalgae offering potential for more complete closed-loop systems [11]. Similarly, in terrestrial applications such as offices, schools, and healthcare facilities, algae-based air purifiers can reduce CO2 by 150-300 ppm while increasing oxygen levels by 2-4% without elevating indoor humidity to uncomfortable levels [95].

Experimental Workflow Comparison: This diagram contrasts the controlled processes of microalgae cultivation with the more variable growth conditions of higher plants, highlighting differences in oxygen production and humidity outcomes.

The comparative analysis of oxygen production and humidity impact reveals distinct performance advantages of microalgae systems over higher plants for engineered air revitalization applications. Microalgae provide superior oxygen production rates, significantly higher CO2 sequestration efficiency, and critical humidity management capabilities through closed-system cultivation. These characteristics make microalgae particularly suitable for environments requiring precise atmospheric control, including space stations, specialized research facilities, and modern energy-efficient buildings with limited ventilation. Future research directions should focus on optimizing photobioreactor designs, enhancing light delivery systems, and developing more robust algal strains capable of maintaining high productivity under varying environmental conditions. The integration of microalgae-based air revitalization represents a promising bio-technological solution for sustainable atmospheric management in confined environments.

The escalating challenges of air pollution and climate change have intensified the search for effective air revitalization technologies. Within this context, biological air purification, which uses living organisms to fix pollutants, presents a sustainable pathway. This guide provides a objective comparison between two principal biological systems: conventional air purifiers (representing physico-chemical technologies) and emerging microalgae-based air purification technologies (MAPT). The analysis is framed within a broader thesis on microalgae versus higher plants for air revitalization efficiency, examining both techno-economic feasibility and environmental impact through Life Cycle Assessment (LCA). Microalgae, as photosynthetic microorganisms, offer a paradigm shift by metabolizing pollutants into biomass, contrasting with the filter-based capture of conventional systems and the slower remediation rates of terrestrial plants [43] [25].

The following tables synthesize key experimental and commercial data for a direct comparison of the two technologies.

Table 1: Techno-Economic Performance Comparison

Performance Parameter	Conventional Air Purifiers	Microalgae-Based Air Purifiers (MAPT)
Primary Purification Mechanism	Adsorption/Filtration on synthetic materials [43]	Biological fixation via photosynthesis [43]
CO2 Fixation Efficiency	Not applicable; may be a source	10–50 times higher than terrestrial plants [43] [7]
Typical CO2 Sequestration	-	~1.3-1.7 kg CO2 per kg biomass [25] [7]
Oxygen Production	None	O2-rich air as a byproduct [43]
Capital Cost	Low to moderate [43]	High (cultivation system costs) [43]
Operational Cost	Moderate (periodic filter replacement) [43]	Moderate (culture maintenance, harvesting) [39]
By-product Generation	Spent filters (hazardous waste) [43]	Valuable biomass for biofuels, feed, fertilizers [43] [99] [100]

Table 2: Life Cycle Environmental Impact Assessment (LCA) Summary

LCA Impact Category	Conventional Air Purifiers	Microalgae-Based Air Purifiers	Notes
Global Warming Potential	Higher (linked to grid electricity use) [100]	Lower, especially with renewable energy [100]	Fish feed product shows lowest impact [100].
Resource Depletion	Higher (consumable filters, materials) [43]	Lower (utilizes CO2 and sunlight) [43]
Waste Generation	Significant (non-recyclable filters) [43]	Minimal to zero waste in biorefinery model [43] [99]	MAPT biomass can be fully valorized.
Overall LCA Impact	Higher negative impact per functional unit [43]	Lower negative LCA impact, but data is incomplete [43]	LCA is highly sensitive to energy source and system design.

Experimental Protocols for Performance Evaluation

To ensure the comparability and reliability of data cited in this guide, the following outlines the standard experimental methodologies employed in the field.

Protocol for Microalgae Cultivation and Air Purification Trials

This protocol describes the setup for evaluating the air revitalization performance of microalgae in controlled photobioreactors [43] [39].

• Strain Selection and Inoculation: Select target microalgae species (e.g., Chlorella vulgaris). Inoculate a sterile BG-11 medium with the pre-cultured algal strain to an initial optical density (OD680) of 0.1 [39].
• Photobioreactor Setup: Use a closed photobioreactor system, such as an airlift photobioreactor (AL-PBR). Maintain temperature at 25 ± 1°C. Provide continuous illumination with cool white fluorescent lamps at a specified light intensity (e.g., 120 μmol m⁻² s⁻¹) [39].
• Pollutant Gas Supply: Continuously sparge the culture with a simulated polluted air stream. A standard mixture contains 5% CO2 (v/v) in air, supplied at a defined flow rate (e.g., 0.1 vvm - gas volume per liquid volume per minute) [39].
• Monitoring and Analytics:
  • Growth Kinetics: Monitor algal growth daily by measuring optical density and dry cell weight.
  • Gas Analysis: Use a gas analyzer to measure the concentration of CO2 at the inlet and outlet of the PBR to calculate fixation rates [43].
  • Pollutant Removal: Periodically sample the culture and effluent gas to quantify the removal efficiency of target pollutants (e.g., NOx, VOCs) via standardized analytical methods like GC-MS or HPLC [43].
• Biomass Harvesting: At the end of the batch or in a semi-continuous mode, harvest biomass. A common method involves flocculation using a biodegradable flocculant like cationic starch, followed by filtration or gravity sedimentation [39].

Protocol for Life Cycle Assessment (LCA) of Air Purification Systems

LCA is a standardized methodology (ISO 14040/14044) used to evaluate the comprehensive environmental impacts of a product system from raw material extraction to end-of-life ("cradle-to-grave") [100] [101].

• Goal and Scope Definition: Define the purpose and the functional unit (e.g., "treatment of 1000 m³ of indoor air to WHO standards"). Establish the system boundaries to include all relevant processes (e.g., for MAPT: culture construction, nutrient production, electricity for mixing/lighting, harvesting, and processing) [100] [101].
• Life Cycle Inventory (LCI): Compile and quantify all material and energy inputs (water, nutrients, electricity, CO2) and environmental outputs (emissions to air, water, soil) for each process within the system boundaries. This relies on industrial process data and specialized LCA databases [100].
• Life Cycle Impact Assessment (LCIA): Translate the LCI data into potential environmental impact categories using characterization models. Common categories include Global Warming Potential, Freshwater Ecotoxicity, Human Carcinogenic Toxicity, and Water Consumption [100].
• Interpretation: Analyze the results to identify significant contributors to environmental impacts, assess sensitivity to key parameters (e.g., source of electricity), and draw conclusions and recommendations for environmental performance improvement [43] [100].

System Workflows and Pathway Analysis

The logical relationship and material flows of the two contrasted systems can be visualized as follows.

Air Purification System Comparison

Microalgae Biorefinery Value Chain

The pathway for valorizing harvested microalgae biomass into multiple products, a key economic advantage, is detailed below.

The Scientist's Toolkit: Key Research Reagents and Materials

Table 3: Essential Materials and Reagents for Microalgae Air Purification Research

Item	Function in Research Context
Model Microalgae Strains (e.g., Chlorella vulgaris, Spirulina platensis)	Model organisms for studying photosynthetic gas exchange, pollutant fixation efficiency, and biomass productivity under controlled conditions [39] [102].
Photobioreactor (PBR) Systems	Controlled environments (closed PBRs or open raceway ponds) for cultivating microalgae and precisely monitoring gas-liquid mass transfer and purification performance [43] [25].
Cationic Starch / Chitosan	Biodegradable, non-toxic flocculants used in the harvesting stage to aggregate microalgae cells, facilitating separation from the culture medium [39].
BG-11 / Bold's Basal Medium	Standardized nutrient media providing essential macronutrients (Nitrogen, Phosphorus) and micronutrients for optimal microalgae growth [39].
Gas Analyzer	Instrumentation for real-time measurement of inlet and outlet gas concentrations (e.g., CO2, O2) to quantify gas fixation and exchange rates [43].
Ionic Liquids	Advanced, tunable solvents used in "green" extraction methodologies to efficiently recover lipids and other valuable bioactive compounds from algal biomass [102].

Conclusion

The comparative analysis conclusively demonstrates that microalgae hold a significant edge over higher plants in air revitalization efficiency, primarily due to their superior photosynthetic rates, 10–50 times greater carbon sequestration capacity, and proven efficacy in removing a wider spectrum of pollutants including PM2.5, PM10, and VOCs. For biomedical research and clinical environments, where air quality is paramount, microalgae-based systems offer a promising, active purification technology. Future research should prioritize the domestication of high-performance algal strains, the development of cost-effective hybrid photobioreactors, and rigorous clinical trials to validate the health benefits of algae-mediated air purification in sensitive settings such as hospitals and laboratories, ultimately paving the way for their integration into advanced environmental control systems.

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