\r\n\t \r\n\tRecently in 2019, International Council on Systems Engineering (INCOSE) has released the latest version of the “Guidelines for the Utilization of ISO/IEC/IEEE 15288 in the Context of System of Systems (SoS) Engineering” to industry for review and comments. The document was developed under the Partner Standards Development Organization cooperation agreement between ISO and IEEE, as it was approved by Council Resolution 49/2007. This document provides guidance for the utilization of ISO/IEC/IEEE 15288 in the context of SoS in many domains, including healthcare, transportation, energy, defense, corporations, cities, and governments. This document treats an SoS as a system whose elements are managerially and/or operationally independent systems, and which together usually produce results that cannot be achieved by the individual systems alone. This INCOSE guide book perceives that SoS engineering demands a balance between linear procedural procedures for systematic activity and holistic nonlinear procedures due to additional complexity from SoS perspectives. \r\n\tThe objective of this book is to provide a comprehensive reference on Systems-of-Systems Engineering, Modeling, Simulation and Analysis (MS&A) for engineers and researchers in both system engineering and advanced mathematical modeling fields. \r\n\tThe book is organized in two parts, namely Part I and Part II. Part I presents an overview of SOS, SOS Engineering, SOS Enterprise Architecture (SOSEA) and SOS Enterprise (SOSE) Concept of Operations (CONOPS). Part II discusses SOSE MS&A approaches for assessing SOS Enterprise CONOPS (SOSE-CONOPS) and characterizing SOSE performance behavior. Part II focuses on advanced mathematical application concepts to address future complex space SOS challenges that require interdisciplinary research involving game theory, probability and statistics, non-linear programming and mathematical modeling components.
\r\n
\r\n\tPart I should include topics related to the following areas: \r\n\t- SOS and SOS Engineering Introduction \r\n\t- Taxonomy of SOS \r\n\t- SOS Enterprise (SOSE), SOSE CONOPS, Architecture Frameworks and Decision Support Tools
\r\n
\r\n\tPart II should address the following research areas: \r\n\t- SOS Modeling, Simulation & Analysis (SOS M&SA) Methods \r\n\t- SOS Enterprise Architecture Design Frameworks and Decision Support Tools \r\n\t- SOS Enterprise CONOPS Assessment Frameworks and Decision Support Tools.
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Nguyen received his M.A. in Mathematics, and his Ph.D. in Applied Mathematics from the Claremont Graduate University; M.S.E.E. in Communication Systems Theory from University of California San Diego; and B.S.E. in Electronics and M.S.E. in Electromagnetic Field Theory from California State University Fullerton (CSUF). He also completed all course requirements and passed the comprehensive exam for his M.S.E.E. in Digital Signal Processing from California State University Long Beach. Dr. Nguyen is an expert in Satellite Operations (SATOPS), Satellite Communications (SATCOMs), advanced mathematical modeling for complex systems-of-systems, sensing and communication networks.\nCurrently, he serves as Adjunct Research Professor at CSUF, Mathematics Dept. Concurrently, he is also with the Aerospace Corporation, serving as a Deputy Chief System Architect in Space Systems Architect, Global Partnerships Subdivision. He has more than 13-years of service at Aerospace, and prior to his current position; he has served as Sr. Engineering Specialist, Sr. Project Lead, Section Manager, Associate Director, Interim Director, and Principal Technical Staff (the highest technical level at the corporation). At Aerospace, he invented HPA linearizer, GMSK synchronizers and developed advanced optimization techniques using game theory for achieving affordable and low-risk acquisition strategy. Prior to CSUF, he had also held a Research Assistant Professor at the Catholic University of America in concurrent with The Aerospace Corporation positions. \nHe was a Engineering Fellow from Raytheon, where he had 10-year of services at Raytheon, serving as Program Area Chief Engineer, Program Chief Engineer, PI, Technical Director, Program Manager, Lead Architect and Lead System Engineer for many advanced programs and pursuits related to sensing and communication networks. At Raytheon, he invented radar-communication technology and gun barrel detector using millimeter-wave. Previous to Raytheon and Aerospace Corporation, Dr. Nguyen was with NASA/JPL for more than 11-years, where he served as the NASA delegate to the international Consultative Committee for Space Data System (CCSDS). Many of his works on RF and Modulation were adopted as the CCSDS standards for USB waveforms and space RF systems. At JPL he invented QPSK phase ambiguity resolver and developed innovative optimization technique for simultaneous range-command-telemetry operation. He built the first laser lab and automated manufacturing lab when he was with ITT Technical Services in the early ’80s. \nHe has published more than 250 technical reports and papers. His work has appeared in NASA TechBrief, textbook, Open Access Book, SIAM Publication, CCSDS Blue Book, and Wiley & Sons Encyclopedia of Electrical and Electronics Engineering. He was selected as a Vietnamese-American Role Model by KCSI-TV, Channel 18 in 2002, and Recognition Honoree at 50-Year Celebration of CSUF in 2007. He received numerous Raytheon, Aerospace and NASA awards, and Air Force commendations. He holds 16 patents and has 01 patent pending. 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1. Introduction
The phytoplankton community of open oligotrophic oceans is dominated by prokaryotic Prochlorococcus spp., Synechococcus spp., and eukaryotic pico- and nanophytoplankton [1-3]. The competitive success of these phytoplankton species depends on different factors, including the response to the (dynamic) irradiance conditions encountered in the water column. With the occurrence of different ecotypes, picophytoplankton species such as Prochlorococcus spp., Synechococcus spp., and Ostreococcus spp. can grow over a broad range of irradiance conditions [4-7]. For example, the low light adapted ecotypes of Prochlorococcus are well adapted to the irradiance intensity and spectral composition of the deep chlorophyll maximum with high chlorophyll b/a ratios and low optimal growth irradiances [4,5,8]. In contrast, the high light adapted ecotypes of Prochlorococcus spp. can competitively grow in the (upper) mixed layer with low chlorophyll b/a ratios and higher optimal growth irradiances [4,5,8]. Similar differences in pigmentation, absorption, and photosynthetic characteristics have been found in ecotypes of marine Synechococcus spp. [9-11] and Ostreococcus spp. [7,12,13]. In addition to the genetically defined (photo)physiology of the different ecotypes, the photoacclimation potential of specific (pico)phytoplankton species may play an important role in the response to (dynamic) irradiance conditions [11].
Phytoplankton irradiance exposure is strongly influenced by physical processes in the ocean [14]. During stratification, phytoplankton can be trapped in a shallow upper mixed layer, thereby enhancing exposure to photosynthetically active radiation (PAR, 400-700 nm) and ultraviolet radiation (UVR, 280-400 nm), or can experience limiting irradiance conditions at the deep chlorophyll maximum. In seasonally stratified regions, the period of stratification is interchanged with periods of deep convective mixing that can reach below the euphotic zone. This causes a strong reduction in the daily experienced irradiance, with occasional interruptions of excessive irradiance exposure. Consequently, phytoplankton irradiance exposure in open ocean ecosystems can vary by several orders of magnitude on a time scale ranging from seconds to days. Moreover, short wavelength solar radiation (UVB, 280-315 nm) can penetrate to significant depths in clear oligotrophic waters [15,16].
High irradiance exposure may have considerable effects on photosynthesis and viability in oceanic picophytoplankton species such as Prochlorococcus spp., Synechococcus spp., and Ostreococcus spp. [17-19]. When residing near the surface, picophytoplankton can experience irradiance intensities that exceed photosynthetic requirements. Exposure to excessive PAR and UVR causes photoinhibition, a process in which an over-reduction of the photosynthetic electron transport chain reduces photosynthetic efficiency by a decrease in functional photosystem II (PSII) reaction centers [20]. Moreover, prolonged exposure to excessive irradiance can lead to the uncontrolled formation of reactive oxygen species and viability loss [21,22]. To prevent photoinhibition and viability loss during excessive irradiance exposure, phytoplankton regulate light harvesting and other photosynthetically important processes. In prokaryotic species, the utilization of light harvesting energy can be regulated by state transitions, in which the light harvesting antenna of the phycobillisome (PBS) is redistributed between the reaction centers of photosystem I (PSI) and PSII [23,24]. In addition, light harvesting energy can be regulated by the thermal dissipation of excess energy. This photoprotective process can occur within seconds after irradiance changes in both prokaryotic and eukaryotic phytoplankton species, but the underlying mechanisms are considerably different. In eukaryotic species, the thermal dissipation of excess energy involves the xanthophyll pigment cycle. Epoxidized xanthophyll cycle pigments assist in light harvesting, whereas de-epoxidized equivalents dissipate excess energy in the form of heat [25]. In PBS containing cyanobacteria, the thermal dissipation of excess energy involves the orange carotenoid protein [24,26]. In Prochlorococcus spp., these proteins are not observed and the underlying mechanism remains unknown [24,27]. In addition to the regulation of light harvesting, photoinhibition and viability loss may be avoided by the increase of photochemical quenching by enhancing alternative electron transport and (non-)enzymatic scavenging of reactive oxygen species [28,29]. Simultaneously, phytoplankton can counteract the effects of photoinhibition by photorepair, a process in which damaged D1 proteins are removed from PSII and replaced by newly synthesized D1 proteins [20].
Although it has previously been reported that temperature may have a positive effect on the survival of picophytoplankton under high irradiance conditions [30], no direct assessment of the temperature-dependency of photoregulation during high PAR and UVR exposure is available for this specific phytoplankton group. A recent study showed that both prokaryotic and eukaryotic picophytoplankton may be less susceptible to the negative effects of high irradiance intensities at elevated temperatures [31]. In the prokaryotic species Prochlorococcus spp. (eMED4 and eMIT9313) and the eukaryotic species Ostreococcus sp. (clade B) and Pelagomonas calceolata, acclimation to elevated temperatures enhanced photoacclimation to higher irradiance intensities and reduced photoinhibition [31]. This has also been found in larger phytoplankton species, such as the diatom species Chaetoceros gracilis, Thalassiosira pseudonana, and Thalassiosira weissflogii [32-34]. In cyanobacteria and eukaryotic nanophytoplankton, reduced levels of photoinhibition at elevated temperatures may be associated with enhanced rates of state transitions [24], enhanced enzymatic conversions of the xanthophyll pigment cycle [35], enhanced D1 repair [36], and the potential enhancement of Rubisco activity [34]. However, the potential role of these photoregulating mechanisms at elevated temperatures remains unknown in oceanic picophytoplankton.
In the present study, a comparative analysis of the high irradiance sensitivity of oceanic picophytoplankton was performed to study the combined effect of elevated temperatures and irradiance levels near the surface of open oligotrophic oceans. To this end, two prokaryotic and two eukaryotic strains were acclimated to 16 °C, 20 °C, and 24 °C, after which they were exposed to a single high PAR dose, with and without UVR. The response to and the recovery after high irradiance exposure was assessed by analysis of PSII fluorescence and pigmentation in order to investigate immediate photoinhibition and photoprotective processes. The results are discussed in the context of differences between oceanic picophytoplankton species and are used to unravel the importance of photoinhibition in structuring the phytoplankton community in open oligotrophic oceans.
2. Method
2.1. Culture conditions
Cultures were obtained from the Roscoff Culture Collection (RCC) and the Provasoli-Guillard National Center for Marine Algae and Microbiota (NCMA). The strains were all isolated from oligotrophic regions and are representative for low light (LL) and high light (HL) adapted species in open ocean ecosystems. Prochlorococcus marinus strain CCMP2389 (ecotype MED4, HL) and Prochlorococcus sp. strain RCC407 (ecotype MIT9313, LL) were cultured in K/10-Cu medium based on natural oceanic seawater as described by [37]. Ostreococcus sp. strain RCC410 (clade B, LL) and Pelagomonas calceolata strain RCC879 (LL) were cultured in K medium as described by [38]. Cultures were maintained in 100 ml glass Erlenmeyer flasks at 9 μmol photons m-2 s-1 (Prochlorococcus sp. and P. calceolata) and 68 μmol photons m-2 s-1 (P. marinus and Ostreococcus sp.) in a diurnal cycle of 12:12 h light:dark at 20 °C.
2.2. Experimental design
Cultures of P. marinus, Prochlorococcus sp., Ostreococcus sp., and P. calceolata were transferred to 500 ml glass Erlenmeyer flasks and incubated in triplicate at 16 °C, 20 °C, and 24 °C. Experiments were carried out in a temperature controlled U-shaped lamp setup as described by [39]. The temperature in the setup was maintained at 16 °C, 20 °C, and 24 °C by a thermostat (RK 8 KS, edition 2000, Lauda Dr. R. Wobser & Co.) and deviated less than ± 0.5 °C. During the experiments, 50 μmol photons m-2 s-1 PAR (Biolux and Skywhite lamps, Osram) was provided as a square wave function with a 12:12 h light:dark cycle (monitored with a QSL-100, Biospherical Instruments). Prior to the experiments, the picophytoplankton strains were kept in exponential growth phase and acclimated to the experimental irradiance and temperature conditions for at least three weeks. In mid-exponential growth phase, the response to high photosynthetically active radiation (PAR, 400-700 nm), with and without ultraviolet radiation (UVR, 290-400 nm), was assessed at growth temperature by pigment and PSII chlorophyll fluorescence analysis. To this end, 200 ml of each replicate culture was exposed to high PAR and PAR+UVR for 10 min in a temperature controlled (RTE-211, Neslab Instruments Inc.) irradiance set-up at 16 °C, 20 °C, or 24 °C. The irradiance set-up provided ± 500 µmol photons m-2 s-1 by a 250 W MHN-TD lamp (Philips) and two 20 W TL/12 UVB fluorescent lamps (Philips), in which the PAR and PAR+UVR conditions were obtained by using the long pass filters GG395 and WG305 (Schott AG, Mainz), respectively (Table 1). Prior to exposure (t = 0), samples for the analysis of pigmentation and the maximum quantum yield of PSII (Fv/Fm) were collected and measured as described below. After exposure, treated culture samples were transferred to dim light conditions at growth temperature (16 °C, 20 °C, or 24 °C). Subsequently, samples for the analysis of pigmentation were taken at t = 10, 20, and 40 min and recovery of the quantum yield of PSII (ФPSII) was determined at t = 10, 15, 20, 25, 30, 35, 40, 60, 80, and 100 min for both PAR and PAR+UVR treated cultures. Culturing of Prochlorococcus sp. at 16 °C and 50 µmol photons m-2 s-1 was attempted several times, but this condition exceeded the limit for growth of this individual strain. No measurements were performed for Prochlorococcus sp. under these conditions.
\n\t\t
\n\t\t
\n\t\t
\n\t\t
\n\t\t\t
\n\t\t\t
\n\t\t\t\tPAR\n\t\t\t
\n\t\t\t
\n\t\t\t\tPAR+UVR\n\t\t\t
\n\t\t
\n\t\t
\n\t\t\t
PAR
\n\t\t\t
172
\n\t\t\t
157
\n\t\t
\n\t\t
\n\t\t\t
UVA
\n\t\t\t
8.69
\n\t\t\t
15.3
\n\t\t
\n\t\t
\n\t\t\t
UVB
\n\t\t\t
0.05
\n\t\t\t
1.79
\n\t\t
\n\t
Table 1.
Doses (W m-2) for photosynthetically active radiation (PAR, 400-700 nm) and ultraviolet radiation A (UVA, 315-400 nm) and B (UVB, 290-315 nm) are given for the PAR and PAR+UVR treatments during the experiments. Total irradiance intensity was ± 500 µmol photons m-2 s-1 in both treatments.
2.3. Photosystem II chlorophyll fluorescence characteristics
PSII fluorescence analyses were performed on a WATER-PAM chlorophyll fluorometer (Waltz GmbH) equipped with a WATER-FT flow-through emitter-detector unit and analyzed using WinControl software (version 2.08, Waltz GmbH) according to [40] and references therein. Prior to exposure to PAR and PAR+UVR (t = 0), 5-15 ml culture samples were dark-adapted for 20 min at 16 °C, 20 °C, or 24 °C. For analysis, the measuring light was turned on and F0 was recorded as the minimal fluorescence. During a saturating light flash, Fm° was then recorded as the maximum fluorescence in the dark-adapted state. The maximum quantum yield of PSII (Fv/Fm) was calculated as (Fm° - F0) / Fm°. After exposure (t = 10-100), the quantum yield of PSII (ΦPSII) was determined by measuring Ft as the steady state fluorescence prior to the saturating light flash and Fm’ as the maximum fluorescence in the light. ΦPSII was calculated as (Fm’ – Ft) / Fm’. From the Fv/Fm measurements at t = 0 and the ΦPSII measurements at t = 10, total non photochemical quenching (NPQ) was calculated as (Fm° - Fm’) / Fm’. Relaxation analysis was performed to estimate the contribution of slowly and rapidly relaxing non photochemical quenching. Relaxation of NPQ on a time scale of minutes is associated with photoprotective processes such as state transitions, relaxation of the xanthophyll pigment cycle or other forms of thermal dissipation [35,40,41]. Processes that relax over a longer period of time (hours) are referred to as photoinhibition, i.e. damage to the reaction centers of PSII [40,42]. To estimate photoprotection and photoinhibition, the recorded Fm’ was corrected for baseline quenching by subtracting F0 and was log transformed for further analysis. Transformed Fm’ values of the final 60 min of the ФPSII recovery curve were extrapolated to calculate the value of Fm‘ that would had been attained if only slowly relaxing quenching was present in the light (Fmr). Slow relaxing non photochemical quenching (NPQS) was then calculated as (Fm° - Fmr) / Fmr and fast relaxing non photochemical quenching (NPQF) as (Fm° / Fm’) - (Fm° - Fmr). In addition, the contribution of UVR to the decrease in quantum yield of PSII during irradiance exposure was calculated as (ΦPSII,PAR - ΦPSII,PAR+UVR ) / ΦPSII,PAR * 100 [43].
2.4. Pigment composition
Samples (25-30 ml) for untreated (t = 0), PAR treated (t = 10, 20, 40), and PAR+UVR (t = 10, 20, 40) treated cultures were filtered onto 25 mm GF/F filters (Whatman), snap frozen in liquid nitrogen, and stored at -80 °C until further analysis. Pigments were quantified using High Performance Liquid Chromatography (HPLC) as described by [44]. In short, filters were freeze-dried for 48 h and pigments were extracted in 3 ml 90% acetone (v/v, 48 h, 4 °C). Detection of pigments was carried out using a HPLC (Waters 2695 separation module, 996 photodiode array detector) equipped with a Zorbax Eclipse XDB-C8 3.5 µm column (Agilent Technologies, Inc.). Peaks were identified by retention time and diode array spectroscopy. Pigments were quantified using standards (DHI LAB products) of chlorophyll a1, chlorophyll a2, diadinoxanthin (Dd), diatoxanthin (Dt), violaxanthin (Vio), antheraxanthin (Ant), and zeaxanthin (Zea). From here on, chlorophyll a (Chl-a) will refer to chlorophyll a2 in P. marinus and Prochlorococcus sp. and to chlorophyll a1 in Ostreococcus sp. and P. calceolata. The de-epoxidation state (DPS) of the xanthophyll pigment cycle was calculated as (Ant + Zea) / (Vio + Ant + Zea) for Ostreococcus sp. and as Dt / (Dd + Dt) for P. calceolata. In addition to the DPS, the rate of de-epoxidation of the xanthophyll pigment cycle (kDPS in min-1) was estimated as the increase in DPS during exposure to high PAR and PAR+UVR [45].
2.5. Statistical analysis
All measurements were performed for triplicate cultures (n = 3) at each temperature. Differences between the three temperature conditions, differences between irradiance treatments, and differences between species were statistically tested by analysis of variance (ANOVA) using STATISTICA software (version 8.0 and 10.0, StatSoft Inc.). Before analysis, data were tested for normality and homogeneity of variances. Differences were considered significant when p < 0.05.
3. Results
3.1. Non photochemical quenching and photosystem II recovery
P. marinus, Prochlorococcus sp., Ostreococcus sp., and P. calceolata all showed non photochemical quenching (NPQ) upon exposure to high photosynthetically active radiation (PAR), with and without ultraviolet radiation (UVR) (Figure 1). The effect of temperature on NPQ was most pronounced in the prokaryotic strains P. marinus and Prochlorococcus sp. (Figure 1). Although total NPQ did not change with temperature in P. marinus, the proportion of slow and fast non photochemical quenching changed significantly. Slow relaxing non photochemical quenching (NPQS) decreased with increasing temperature (p < 0.05, not significant between 20 °C and 24 °C), whereas fast relaxing non photochemical quenching (NPQF) increased significantly with increasing temperature (p < 0.05). In Prochlorococcus sp., total NPQ increased from 20 °C to 24 °C (Figure 1). The proportion of NPQS and NPQF was also affected by temperature in Prochlorococcus sp., with a significant increase in NPQF with increasing temperature (p < 0.05) and unchanged levels of NPQS. In the eukaryotic species Ostreococcus sp., temperature had no effect on NPQ (Figure 1). In P. calceolata, total NPQ decreased with increasing temperature (p < 0.05, not significant between 20 °C and 24 °C). This was associated with a decrease in NPQS with increasing temperature (p < 0.05, not significant between 16 °C and 20 °C), whereas NPQF remained unaffected by temperature.
Figure 1.
Non photochemical quenching. Mean (± standard deviation, n = 3) total non photochemical quenching (NPQ), slow relaxing NPQ (NPQS), and fast relaxing NPQ (NPQF) are given for Prochlorococcus marinus eMED4, Prochlorococcus sp. eMIT9313, Ostreococcus sp. clade B, and Pelagomonas calceolata at 16 °C, 20 °C and 24 °C. The picophytoplankton strains were exposed to high photosynthetically active radiation (PAR, white bars) and high PAR with ultraviolet radiation (PAR+UVR, grey bars) for 10 minutes. Significant effects (p < 0.05) of the growth temperature (*) and the spectral composition of the irradiance treatment (“) are indicated.
The spectral composition of the irradiance treatment influenced non photochemical quenching and the recovery of the quantum yield of PSII (ΦPSII) considerably in the prokaryotic strains (Figure 1, Figure 2). In both P. marinus and Prochlorococcus sp., total NPQ and NPQS were significantly higher during exposure to PAR+UVR compared with PAR, whereas NPQF decreased significantly during exposure to UVR (p < 0.05) (Figure 1). In Prochlorococcus sp., this was associated with almost no recovery of ΦPSII after exposure to PAR+UVR (Figure 2). In the eukaryotic species Ostreococcus sp., the spectral composition of the irradiance treatment did not have a significant effect on NPQ (Figure 1). However, recovery of ΦPSII in Ostreococcus sp. was lower after exposure to PAR+UVR compared with PAR (significant for t = 60-100, p < 0.05, Figure 2). In P. calceolata, exposure to PAR+UVR significantly increased NPQS and decreased NPQF (p < 0.05), but total NPQ remained unaffected by the spectral composition of the irradiance treatment (Figure 1). P. calceolata showed no recovery of ΦPSII after exposure to PAR+UVR (Figure 2).
Figure 2.
Recovery of PSII after high irradiance exposure. Mean (± standard deviation, n = 3) quantum yield of PSII (ΦPSII in % of Fv/Fm) during and after exposure to high irradiance for Prochlorococcus marinus eMED4, Prochlorococcus sp. eMIT9313, Ostreococcus sp. clade B, and Pelagomonas calceolata acclimated to 20 °C. The picophytoplankton strains were exposed to high photosynthetically active radiation (PAR, white circles) and high PAR with ultraviolet radiation (PAR+UVR, dark grey circles) for 10 minutes (light grey area).
Comparison of NPQ between the different picophytoplankton strains demonstrated significantly lower total NPQ in the prokaryotic species P. marinus and Prochlorococcus sp. compared with the eukaryotic species Ostreococcus sp. and P. calceolata (p < 0.05) (Figure 1). In P. calceolata, NPQS was significantly higher compared with the other species (p < 0.05, not significant at 24 °C). P. marinus and Prochlorococcus sp. showed intermediate levels of NPQS, whereas Ostreococcus sp. showed significantly lowest NPQS (p < 0.05, not significant at 24 °C). The relative low levels of NPQS in Ostreococcus sp. were accompanied by significantly higher NPQF compared with the other species (p < 0.005). No differences in NPQF were found between P. marinus, Prochlorococcus sp., and P. calceolata.
3.2. Inhibition of photosystem II by ultraviolet radiation
The inhibition of ΦPSII due to UVR was affected by temperature in P. marinus, Ostreococcus sp., and P. calceolata (Table 2). In P. marinus, UVR inhibition decreased significantly with increasing temperature (p < 0.01 for 16 °C compared with 24 °C). In the eukaryotic species Ostreococcus sp. (not between 20 °C and 24 °C) and P. calceolata, UVR inhibition of ΦPSII also decreased with increasing temperature, but not significantly. In Prochlorococcus sp., no effect of temperature was found on the UVR inhibition of ΦPSII. Comparison of the different picophytoplankton strains showed that Ostreococcus sp. was least inhibited by UVR (p < 0.001) (Figure 2, Table 2). P. marinus showed intermediate levels of UVR inhibition, whereas ΦPSII was most inhibited by UVR in Prochlorococcus sp. and P. calceolata (p < 0.001).
Mean (± standard deviations, n = 3) inhibition by ultraviolet radiation (% of photosynthetically active radiation treatment) after 10 min high irradiance exposure in Prochlorococcus marinus eMED4, Prochlorococcus sp. eMIT9313, Ostreococcus sp. clade B, and Pelagomonas calceolata acclimated to 16 °C, 20 °C, and 24 °C. abc indicate significant effects of the temperature treatment within each species. n/a: data not available, growth was not observed under the used conditions and no additional measurements were performed.
3.3. Photoprotective pigmentation
Temperature acclimation affected the initial photoprotective pigment pool in P. marinus (t = 0, Table 3), with higher zeaxanthin per chlorophyll a levels at lower temperatures (p < 0.001). In Prochlorococcus sp., no significant effect of temperature acclimation was observed in the initial zeaxanthin per chlorophyll a level. In both prokaryotic strains, exposure to high irradiance did not influence photoprotective pigmentation, as the zeaxanthin per chlorophyll a levels remained similar during and after high irradiance exposure (Table 3). In Ostreococcus sp., acclimation to higher temperatures increased the initial xanthophyll cycle pigment pool (30-40%), but not significantly (t = 0, Table 3). In response to high irradiance exposure, large fluctuations in the sum of violaxanthin, antheraxanthin, and zeaxanthin per chlorophyll a were observed and no significant effect of temperature acclimation on the photoprotective pigment pool was found (Table 3). In P. calceolata, the initial photoprotective pigments per chlorophyll a ratio was highest at 24 °C (19 %, not significant). Temperature had no effect on the total xanthophyll cycle pigment pool in response to high irradiance in P. calceolata as the sum of diadinoxanthin and diatoxanthin per chlorophyll a remained unchanged during and after exposure to high irradiance (Table 3). No significant effect of the spectral composition of the irradiance treatment was observed in the photoprotective pigments pools of P. marinus, Prochlorococcus sp., Ostreococcus sp., and P. calceolata (Table 3).
\n\t\t
\n\t\t
\n\t\t
\n\t\t
\n\t\t
\n\t\t
\n\t\t
\n\t\t
\n\t\t\t
\n\t\t\t
\n\t\t\t\tPAR\n\t\t\t
\n\t\t\t
\n\t\t\t\tPAR+UVR\n\t\t\t
\n\t\t
\n\t\t
\n\t\t\t
\n\t\t\t
\n\t\t\t\t16 °C\n\t\t\t
\n\t\t\t
\n\t\t\t\t20 °C\n\t\t\t
\n\t\t\t
\n\t\t\t\t24 °C\n\t\t\t
\n\t\t\t
\n\t\t\t\t16 °C\n\t\t\t
\n\t\t\t
\n\t\t\t\t20 °C\n\t\t\t
\n\t\t\t
\n\t\t\t\t24 °C\n\t\t\t
\n\t\t
\n\t\t
\n\t\t\t
\n\t\t\t\tProchlorococcus marinus\n\t\t\t
\n\t\t
\n\t\t
\n\t\t\t
t = 0
\n\t\t\t
0.647±0.060a\n\t\t\t
\n\t\t\t
0.499±0.004a\n\t\t\t
\n\t\t\t
0.431±0.007a\n\t\t\t
\n\t\t\t
0.647±0.060b\n\t\t\t
\n\t\t\t
0.499±0.004b\n\t\t\t
\n\t\t\t
0.431±0.007b\n\t\t\t
\n\t\t
\n\t\t
\n\t\t\t
t = 10
\n\t\t\t
0.644 ± 0.081
\n\t\t\t
0.488 ± 0.019
\n\t\t\t
0.434 ± 0.017
\n\t\t\t
0.649 ± 0.057
\n\t\t\t
0.488 ± 0.013
\n\t\t\t
0.426 ± 0.017
\n\t\t
\n\t\t
\n\t\t\t
t = 20
\n\t\t\t
0.657 ± 0.066
\n\t\t\t
0.493 ± 0.006
\n\t\t\t
0.426 ± 0.031
\n\t\t\t
0.655 ± 0.071
\n\t\t\t
0.494 ± 0.002
\n\t\t\t
0.421 ± 0.031
\n\t\t
\n\t\t
\n\t\t\t
t = 40
\n\t\t\t
0.661 ± 0.067
\n\t\t\t
0.498 ± 0.009
\n\t\t\t
0.424 ± 0.014
\n\t\t\t
0.663 ± 0.053
\n\t\t\t
0.492 ± 0.013
\n\t\t\t
0.437 ± 0.014
\n\t\t
\n\t\t
\n\t\t\t
\n\t\t\t\tProchlorococcus sp.
\n\t\t
\n\t\t
\n\t\t\t
t = 0
\n\t\t\t
n/a
\n\t\t\t
1.062 ± 0.034
\n\t\t\t
1.025 ± 0.023
\n\t\t\t
n/a
\n\t\t\t
1.062 ± 0.034
\n\t\t\t
1.025 ± 0.023
\n\t\t
\n\t\t
\n\t\t\t
t = 10
\n\t\t\t
n/a
\n\t\t\t
1.206 ± 0.076
\n\t\t\t
0.976 ± 0.009
\n\t\t\t
n/a
\n\t\t\t
1.198 ± 0.039
\n\t\t\t
0.946 ± 0.039
\n\t\t
\n\t\t
\n\t\t\t
t = 20
\n\t\t\t
n/a
\n\t\t\t
1.209 ± 0.093
\n\t\t\t
0.966 ± 0.036
\n\t\t\t
n/a
\n\t\t\t
1.189 ± 0.059
\n\t\t\t
0.936 ± 0.032
\n\t\t
\n\t\t
\n\t\t\t
t = 40
\n\t\t\t
n/a
\n\t\t\t
1.226 ± 0.088
\n\t\t\t
0.996 ± 0.041
\n\t\t\t
n/a
\n\t\t\t
1.192 ± 0.044
\n\t\t\t
0.974 ± 0.039
\n\t\t
\n\t\t
\n\t\t\t
\n\t\t\t\tOstreococcus sp.
\n\t\t
\n\t\t
\n\t\t\t
t = 0
\n\t\t\t
0.079 ± 0.030
\n\t\t\t
0.057 ± 0.024
\n\t\t\t
0.061 ± 0.014
\n\t\t\t
0.079 ± 0.030
\n\t\t\t
0.057 ± 0.024
\n\t\t\t
0.061 ± 0.014
\n\t\t
\n\t\t
\n\t\t\t
t = 10
\n\t\t\t
0.109 ± 0.017
\n\t\t\t
0.062 ± 0.026
\n\t\t\t
0.084 ± 0.032
\n\t\t\t
0.058 ± 0.004
\n\t\t\t
0.053 ± 0.012
\n\t\t\t
0.060 ± 0.013
\n\t\t
\n\t\t
\n\t\t\t
t = 20
\n\t\t\t
0.091 ± 0.036
\n\t\t\t
0.052 ± 0.020
\n\t\t\t
0.060 ± 0.009
\n\t\t\t
0.109 ± 0.003
\n\t\t\t
0.082 ± 0.002
\n\t\t\t
0.105 ± 0.008
\n\t\t
\n\t\t
\n\t\t\t
t = 40
\n\t\t\t
0.106 ± 0.005
\n\t\t\t
0.078 ± 0.006
\n\t\t\t
0.074 ± 0.031
\n\t\t\t
0.109 ± 0.003
\n\t\t\t
0.079 ± 0.014
\n\t\t\t
0.077 ± 0.020
\n\t\t
\n\t\t
\n\t\t\t
\n\t\t\t\tPelagomonas calceolata\n\t\t\t
\n\t\t
\n\t\t
\n\t\t\t
t = 0
\n\t\t\t
0.089 ± 0.008
\n\t\t\t
0.089 ± 0.005
\n\t\t\t
0.106 ± 0.012
\n\t\t\t
0.089 ± 0.008
\n\t\t\t
0.089 ± 0.005
\n\t\t\t
0.106 ± 0.012
\n\t\t
\n\t\t
\n\t\t\t
t = 10
\n\t\t\t
0.096 ± 0.009
\n\t\t\t
0.095 ± 0.002
\n\t\t\t
0.106 ± 0.013
\n\t\t\t
0.092 ± 0.010
\n\t\t\t
0.094 ± 0.004
\n\t\t\t
0.103 ± 0.014
\n\t\t
\n\t\t
\n\t\t\t
t = 20
\n\t\t\t
0.093 ± 0.005
\n\t\t\t
0.096 ± 0.007
\n\t\t\t
0.107 ± 0.014
\n\t\t\t
0.080 ± 0.031
\n\t\t\t
0.093 ± 0.005
\n\t\t\t
0.105 ± 0.014
\n\t\t
\n\t\t
\n\t\t\t
t = 40
\n\t\t\t
0.093 ± 0.009
\n\t\t\t
0.094 ± 0.006
\n\t\t\t
0.109 ± 0.013
\n\t\t\t
0.090 ± 0.008
\n\t\t\t
0.092 ± 0.005
\n\t\t\t
0.104 ± 0.012
\n\t\t
\n\t
Table 3.
Mean (± standard deviations, n = 3) photoprotective pigments per chlorophyll a ratio in Prochlorococcus marinus eMED4 (zeaxanthin), Prochlorococcus sp. eMIT9313 (zeaxanthin), Ostreococcus sp. clade B (violaxanthin, antheraxanthin, and zeaxanthin), and Pelagomonas calceolata (diadinoxanthin and diatoxanthin) acclimated to 16 °C, 20 °C, and 24 °C. Pigment ratios were obtained before (t = 0) and after (t = 10, 20, 40) exposure to high photosynthetically active radiation (PAR), with and without ultraviolet radiation (UVR). abc indicate significant effects of the temperature treatment within each species. n/a: data not available, growth was not observed under the used conditions and no additional measurements were performed.
Figure 3.
De-epoxidation of the xanthophyll pigment cycle. Mean (± standard deviation, n = 3) de-epoxidation state (DPS) of the xanthophyll pigment cycle in Ostreococcus sp. clade B and Pelagomonas calceolata are given during and after 10 minutes of exposure to high photophotosynthetically active radiation (PAR, white circles) and high PAR with ultraviolet radiation (PAR+UVR, grey circles) at 16 °C, 20 °C, and 24 °C.
3.4. De-epoxidation of the xanthophyll cycle
In both Ostreococcus sp. and P. calceolata, the de-epoxidation state (DPS) of the xanthophyll pigment cycle increased significantly during exposure to high irradiance (p < 0.001) (Figure 3). In both strains, the DPS of the xanthophyll pigment cycle decreased over time, but the DPS did not return to initial values after 30 min of recovery in low light conditions (t= 40, Figure 3). In Ostreococcus sp., the de-epoxidation of the xanthophyll pigment cycle mainly included the de-epoxidation of violaxanthin to antheraxanthin, whereas the de-epoxidation of antheraxanthin to zeaxanthin was small. Temperature had an effect on the DPS of the xanthophyll pigment cycle in Ostreococcus sp. (Figure 3, Table 4), but differences were mostly not significant. The initial DPS of the xanthophyll pigment cycle (t = 0) in Ostreococcus sp. was 21-47% higher at 16 °C compared with 20 °C and 24 °C. During exposure to high PAR and PAR+UVR, the increase in the DPS was fastest at 20 °C (Table 4), as was the epoxidation of the xanthophyll pigment cycle after exposure to high irradiance (Figure 3). In P. calceolata, the initial DPS of the xanthophyll pigment cycle was 22-28% lower at 16 °C compared with the higher temperatures (not significant) (Figure 3). During irradiance exposure, the rate of de-epoxidation of the xanthophyll pigment cycle increased with increasing temperature in P. calceolata (not significant) (Figure 3, Table 4). In accordance with the rate of de-epoxidation, the epoxidation of the xanthophyll pigment cycle was fastest at 24 °C (p < 0.05).
The effect of the spectral composition of the irradiance treatment on the de-epoxidation of the xanthophyll pigment cycle was most evident in Ostreococcus sp. (Figure 3). During irradiance exposure (t = 0-10), the DPS in Ostreococcus sp. did not differ significantly between the PAR and PAR+UVR treatment (Figure 3, Table 4). However, in the PAR treatment, epoxidation of the xanthophyll pigment cycle started directly after exposure (t = 10), whereas the epoxidation was delayed in the PAR+UVR treatment and started after 10 minutes of recovery in low light (t = 20). After 30 minutes of recovery (t = 40), the DPS in Ostreococcus sp. was similar in both PAR and PAR+UVR treatments (Figure 3). In P. calceolata, no significant effect of the spectral composition of the irradiance treatment was found, but it seemed that exposure to UVR limited the de-epoxidation of the xanthophyll pigment cycle, especially at lower temperatures (Figure 3).
When the dynamics of the xanthophyll pigment cycle of both species were compared, it was shown that Ostreococcus sp. had a significantly higher DPS compared with P. calceolata (p < 0.05) (Figure 3). In addition, the increase in de-epoxidation of the xanthophyll pigment cycle during high irradiance exposure was faster in Ostreococcus sp. (p < 0.05) (Table 3), whereas no differences in epoxidation rate were observed between Ostreococcus sp. and P. calceolata.
Mean (± standard deviation, n = 3) rate of increase in the de-epoxidation state of the xanthophyll pigment cycle (kDPS in min-1) in Ostreococcus sp. clade B and Pelagomonas calceolata during exposure to high photosynthetically active radiation (PAR) and high PAR with ultraviolet radiation (PAR+UVR) at 16 °C, 20 °C, and 24°C. abc indicate significant effects of the temperature treatment within each species.
4. Discussion
Climate change is expected to mediate a rise in seawater temperature by 1.5-4.5 °C over the next century [46]. This rise in seawater temperature will lead to changes in water column stratification in open oligotrophic oceans [47,48]. The subsequent modifications in mixed layer dynamics increase the exposure of phytoplankton to high levels of photosynthetic active radiation (PAR) and ultraviolet radiation (UVR). Because temperature and irradiance conditions play an important role in the success of specific oceanic phytoplankton species [4,49,50], it is important to understand how oceanic phytoplankton will respond to elevated temperatures and whether this will affect their (photo)physiological performance. The present study focused on the temperature-dependence of photoinhibition and photoregulating processes that are essential for survival during high (dynamic) irradiance conditions.
During short periods of high irradiance exposure, both the prokaryotic picophytoplankton strains P. marinus and Prochlorococcus sp., as the eukaryotic picophytoplankton strains Ostreococcus sp. and P. calceolata were susceptible to photoinhibition. The response to high irradiances was species specific and appeared to be related to the genetically defined light adaptation of the different strains. In the prokaryotic species, the low light adapted ecotype Prochlorococcus sp. (eMIT9313) was highly sensitive to high PAR and UVR, whereas the high light adapted ecotype P. marinus (eMED4) showed lower sensitivity. Similar differences in photoinhibition during high irradiance exposure were observed for other low and high light adapted ecotypes of Prochlorococcus spp. during exposure to high blue irradiance [18]. The differential response to excessive irradiance intensities found in the present study related well to the occurrence of different Prochlorococcus ecotypes in the upper mixed layer (eMED4) and the deep chlorophyll maximum (eMIT9313) [4,49]. In the eukaryotic species, the levels of total non photochemical quenching induced by a tenfold increase in irradiance intensity were similar compared with earlier observations for Ostreococcus sp. and P. calceolata [12,51]. Although the two eukaryotic species were both isolated at 100 m depth from oceanic regions, Ostreococcus sp. showed considerably lower levels of photoinhibition compared with P. calceolata, especially during UVR exposure. It therefore seems that Ostreococcus sp. clade B is not specifically adapted to low light [7], but rather adapted to open ocean irradiance conditions (also see [50,52]) with a relatively low sensitivity to high irradiance intensities compared with other oceanic picophytoplankton [this study,11,31]. The low light adapted ecotype P. calceolata showed highest levels of photoinhibition during exposure to high PAR compared with the other species. However, photoinhibition increased dramatically in the prokaryotic strains during exposure to UVR. This confirms the relative sensitivity of Prochlorococcus spp. to high levels of UVR, as has been observed in oligotrophic waters [53,54].
Temperature acclimation influenced photoinhibition and related processes during high irradiance exposure in P. marinus, Prochlorococcus sp., Ostreococcus sp., and P. calceolata. The effect was not uniform among the different strains, but temperature acclimation influenced the response to high irradiance exposure by changes in the relative contribution of photoinhibition and photoprotective mechanisms to non photochemical quenching in all strains. This general response corresponds well with the observation that both prokaryotic and eukaryotic picophytoplankton may benefit from high irradiance intensities at elevated temperatures by alterations in photophysiology and electron transport [31]. In addition, elevated temperatures had a beneficial effect on the response to high irradiance intensities by partially counteracting the UVR-induced photoinhibition in P. marinus, Ostreococcus sp., and P. calceolata. This was earlier observed in several diatom species and related to an increase in Rubisco activity and gene expression in Thalassiosira weissflogii [34], an increase in repair rates in T. pseudonana [32], and an increase in photoprotection by the dissipation of excess energy in T. weissflogii and C. gracillis [33]. In this study, fast relaxing non photochemical processes, i.e. photoprotection, and the influence of temperature acclimation on these processes was further investigated in the response to excessive irradiance intensities in oceanic picophytoplankton.
Both low and high light adapted Prochlorococcus strains were capable of producing fast relaxing non photochemical quenching (NPQF). Interestingly, the level of NPQF in the low light adapted strain Prochlorococcus sp. (eMIT9313/clade LLIV) was considerably higher compared with that of another low light adapted strain of Prochlorococcus (strain SS120/clade LLII) [27]. It therefore seems that some low light adapted ecotypes of Prochlorococcus are capable of inducing high levels of NPQF comparable to that of high light adapted ecotypes (this study), but others are not [27]. This might possibly be related to the differential occurrence of pcb genes encoding the major chlorophyll binding and light harvesting antenna proteins in both low and high light adapted ecotypes of Prochlorococcus [27,55]. Although the precise underlying mechanism remains unknown, the process of NPQF in P. marinus and Prochlorococcus sp. was sensitive to changes in temperature. It is therefore likely that the underlying mechanisms of NPQF in Prochlorococcus spp. involves an enzymatic reaction or changes due to the improved fluidity of the thylakoid membrane at elevated temperatures [56,57]. This contrasts to earlier observations of NPQF in phycobillisome containing cyanobacteria [58] (for a review see [24]), which supports the notion that the underlying mechanisms are different between Prochlorococcus spp. and other prokaryotic species [24]. It was further shown in the present study that the mechanism of photoprotection in P. marinus and Prochlorococcus sp. was highly sensitive to UVR, possibly related to increased oxidative stress on the thylakoid membrane [59]. Fast relaxing non photochemical quenching was not related to changes in pigmentation during high irradiance exposure in P. marinus and Prochlorococcus sp. The xanthophyll pigment zeaxanthin is not regulated by an epoxydation/de-epoxidation cycle in prokaryotic species and its function is often debated [60,61]. However, the photoprotective role of zeaxanthin is not excluded, since the concentration of zeaxanthin increases relative to chlorophyll a in high light acclimated cells [8,11,61] and zeaxanthin is found in high concentrations in the field [62,63]. The presence of zeaxanthin might have overestimated the calculation of photoinhibition by slowly relaxing non photochemical quenching in the light-harvesting antenna of PSII (F0 quenching) [40,64]. This was however, not observed in P. marinus and Prochlorococcus sp. (data not shown), suggesting that slowly relaxing non photochemical quenching related to damage to the reaction center of PSII in these strains.
In the eukaryotic picophytoplankton species Ostreococcus sp. and P. calceolata, fast relaxing non photochemical quenching coincided with the de-epoxidation of the xanthophyll pigment cycle. The rate of de-epoxidation of the xanthophyll pigment cycle in Ostreococcus sp. and P. calceolata was within the range reported for other eukaryotic pico- and nanophytoplankton [45], as was the relative increase in the de-epoxidation state of the xanthophyll pigment cycle upon high irradiance exposure [12,19,45,51]. For Ostreococcus sp. clade B it was previously shown that both the xanthophyll pigment cycle [19] and alternative electron transport [13] play an important role in the response to high irradiance, whereas photorepair is relatively slow compared with other Ostreococcus ecotypes [19]. This study showed that the photoprotective processes were also effective during UVR exposure, since Ostreococcus sp. was the only strain used in this study that showed substantial NPQF during UVR exposure. The influence of temperature acclimation was also most pronounced during UVR exposure, especially on the xanthophyll pigment cycle. Different effects may add to the high levels of fast relaxing non photochemical quenching observed in Ostreococcus sp. The xanthophyll cycle pigments may have an additional photoprotective function in Ostreococcus sp., including the stabilization of the thylakoid membrane by antheraxanthin and zeaxanthin, providing protection against reactive oxygen species under conditions of a highly reduced electron transport chain (for a review see [65]). In addition, the de-epoxidation of the xanthophyll pigment cycle and the consequent non photochemical quenching in Ostreococcus sp. may be promoted by an increase in the trans-membrane proton gradient due to the presence of chlororespiratory electron flow [13,65]. In P. calceolata, the rate of de-epoxidation and the relative de-epoxidation of the xanthophyll pigment cycle increased at elevated temperature, but this was not associated with an increase in fast relaxing non photochemical quenching. It is possible that the membrane stability necessary for the dissipation of excess energy trough the xanthophyll pigment cycle was affected by oxidative stress [66,67]. This could also explain the diminished fast relaxing non photochemical levels during UVR exposure in this species. Because P. calceolata is a low light adapted ecotype, this species might possibly use additional photoprotective mechanisms, such as the chlororespiratory electron flow observed in Ostreococcus sp., to a lesser extent.
This study showed that oceanic picophytoplankton were susceptible to photoinhibition during short periods of high irradiance. The genetically defined light adaptation of P. marinus, Prochlorococcus sp., Ostreococcus sp., and P. calceolata played an important role in their PAR and UVR sensitivity, likely related to the presence of different (combinations of) photoprotective mechanisms. Temperature acclimation influenced the response to excessive irradiance exposure by changes in the relative contribution of photoinhibition and photoprotective mechanisms to non photochemical quenching. These changes were found to be species specific. Acclimation to elevated temperatures increased the dissipation of excess energy in both P. marinus and Prochlorococcus sp., indicating a strong dependence on temperature of this photoprotective mechanism. In combination with decreased photoinhibition during both PAR and UVR exposure at elevated temperature, the high light adapted ecotype P. marinus may benefit considerably from elevated temperatures in response to high irradiance intensities encountered in the upper mixed layer of open oligotrophic oceans. Considering exposure to UVR, the effect of elevated temperature was most pronounced in the eukaryotic strain Ostreococcus sp., indicating that this species can effectively regulate light harvesting in relatively warm, UVR rich waters near the surface of the open oligotrophic ocean. Even though Prochlorococcus sp. and P. calceolata are unlikely to experience high irradiance intensities in the deep chlorophyll maximum, photoinhibition in these low light adapted ecotypes is highly relevant, since damage to PSII can occur at relatively low irradiance intensities [18,31,68]. At elevated temperatures, the prokaryotic strain Prochlorococcus sp. benefitted by increasing dissipation of excess energy, whereas the eukaryotic strain P. calceolata was less susceptible to photoinhibition. Overall, the differential response to high irradiance may have considerably effect on phytoplankton species distribution and community composition in the open oligotrophic oceans, with some ecotypes and/or species being more susceptible to photoinhibition than others. Photoinhibition and/or photoprotective processes may be positively affected by the rise in seawater temperature associated with climate change, but species specific differences in (photo)physiology remain important in the performance of oceanic picophytoplankton.
Acknowledgments
Remote access to the Roscoff Culture Collection of strains RCC407 and RCC879 was facilitated by ASSEMBLE grant number 227799 (GK). This work was supported by the Netherlands Organization for Scientific Research (NWO), grant numbers 817.01.009 (GK) and 839.08.422 (WHP).
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Introduction ",level:"1"},{id:"sec_2",title:"2. Method",level:"1"},{id:"sec_2_2",title:"2.1. Culture conditions",level:"2"},{id:"sec_3_2",title:"2.2. Experimental design",level:"2"},{id:"sec_4_2",title:"2.3. Photosystem II chlorophyll fluorescence characteristics",level:"2"},{id:"sec_5_2",title:"2.4. Pigment composition",level:"2"},{id:"sec_6_2",title:"2.5. Statistical analysis",level:"2"},{id:"sec_8",title:"3. Results",level:"1"},{id:"sec_8_2",title:"3.1. Non photochemical quenching and photosystem II recovery",level:"2"},{id:"sec_9_2",title:"3.2. Inhibition of photosystem II by ultraviolet radiation",level:"2"},{id:"sec_10_2",title:"3.3. Photoprotective pigmentation",level:"2"},{id:"sec_11_2",title:"3.4. De-epoxidation of the xanthophyll cycle",level:"2"},{id:"sec_13",title:"4. Discussion",level:"1"},{id:"sec_14",title:"Acknowledgments",level:"1"},{id:"sec_14",title:"Acknowledgments",level:"2"}],chapterReferences:[{id:"B1",body:'Li WKW. 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Department of Ocean Ecosystems, Energy and Sustainability Research Institute Groningen, University of Groningen, Groningen, The Netherlands
'},{corresp:null,contributorFullName:"Pablo de Vries",address:null,affiliation:'
Department of Ocean Ecosystems, Energy and Sustainability Research Institute Groningen, University of Groningen, Groningen, The Netherlands
'},{corresp:null,contributorFullName:"Willem H. van de Poll",address:null,affiliation:'
Department of Biological Oceanography, Royal Netherlands Institute for Sea Research, Den Burg, The Netherlands
'},{corresp:null,contributorFullName:"Ronald J. W. Visser",address:null,affiliation:'
Department of Ocean Ecosystems, Energy and Sustainability Research Institute Groningen, University of Groningen, Groningen, The Netherlands
'},{corresp:null,contributorFullName:"Anita G. J. Buma",address:null,affiliation:'
Department of Ocean Ecosystems, Energy and Sustainability Research Institute Groningen, University of Groningen, Groningen, The Netherlands
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\n
1. Introduction
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Frequently referred to as the “the powerhouse of the cell,” the mitochondrion is the key organelle that contributes to neuronal energy and viability through the production of adenosine triphosphate (ATP) via oxidative phosphorylation. During oxidative phosphorylation, electrons from FADH2 or NADH travel across the electron transport chain (ETC) creating an electrical gradient along the inner mitochondrial membrane allowing protons to diffuse through the ATP synthase. This allows the ATP synthase to bind a phosphate group to adenosine diphosphate (ADP) creating ATP. Compared to other metabolic pathways such as fermentation and anaerobic respiration, oxidative phosphorylation is the most efficient process to generate ATP. Energy in the brain is used for overall maintenance of cellular processes, neuronal growth, and axonal branching [1]. However, a majority of the ATP produced is utilized to support one of the neuron’s most essential functions, synaptic transmission [2]. For example, the Na+, K+ -ATPase or the Na+, K+-pump is responsible for approximately half of the energy consumed by the brain, through its use of active transport to pump out sodium ions while taking in potassium ions [3]. This pump is essential in neurotransmission through its regulation of membrane potential, cell volume, and intracellular Ca2+ homeostasis [4, 5]. Likewise, exocytosis requires sufficient energy to release neurotransmitters from presynaptic to postsynaptic vesicles [5].
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Beyond the mitochondria’s role in energy production, it is also a key regulator of apoptotic cell death [6]. Various proteins, such as cytochrome c, reside in the mitochondria and participate in apoptotic pathways. In normal physiological conditions, cytochrome c plays a role as an electron carrier in the ETC. However, during neurotoxic conditions, permeabilization of the mitochondrial membrane occurs, and cytochrome c is released into the cytoplasm. Upon release, cytochrome c binds to apoptotic protease activating factor 1 (Apaf-1) which, in turn, activates caspase-9, forming the apoptosome that then activates downstream caspases leading to cell death [7]. Second mitochondria-derived activator of caspase (Smac)/direct IAP-binding protein with low PI (DIABLO) is also a mitochondrial protein released during apoptosis. The N-terminus of Smac/DIABLO directly interacts with inhibitor of apoptosis proteins (IAPs), a family of proteins that inhibit caspase 3, 7, and 9 activities; thus Smac/ DIABLO exhibits pro-apoptotic roles [8].
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It has been well studied that Bcl-2 family of proteins controls neuronal survival or death via regulating apoptotic pathways, i.e., pro-apoptotic proteins versus anti-apoptotic proteins [9]. The presence of at least one of the four Bcl-2 homology (BH) domains influences a Bcl-2 family member’s role in apoptosis. Pro-apoptotic Bcl-2 family members include the multidomain homology proteins such as Bax and Bak as well as the BH3-only homology proteins such as Bid, Bim, Bad, PUMA, and NOXA. These pro-apoptotic Bcl-2 proteins enhance mitochondrial membrane permeabilization resulting in subsequent release of cytochrome c [10, 11, 12, 13, 14].
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Anti-apoptotic proteins of the Bcl-2 family include Bcl-2, Bcl-xL, Bcl-w, Mcl-1, and Bfl-1. These proteins contain the BH4 homology, which is essential for anti-apoptotic functionality. Both Bcl-2 and Bcl-xL antagonize pro-apoptotic members to prevent apoptosis [15, 16]; for instance, Bcl-xL targets Bak, preventing its oligomerization and inhibiting it from damaging the mitochondrial outer membrane [17]. Similarly, the C-terminal of Bcl-xL binds to the BH3 domain of Bax, resulting in retro-translocation-activated Bax. [18]. Protein-protein binding is further demonstrated with additional members of Bcl-2 family. These observations suggest that the Bcl-2 family’s role in mediating apoptosis and mitochondrial permeabilization is largely influenced by dynamic protein-protein interactions with each other [19, 20, 21].
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The mitochondrion is also responsible for the production of reactive oxygen species (ROS), namely, superoxide and hydrogen peroxide, at Complex I and III of the ETC [22]. This occurs as a result of electron leakage from the complexes, which then allows oxygen to react [23]. Due to the high energy demands required by neuronal mitochondria, this results in increased ROS generation. Increased ROS activity contributes to lipid peroxidation, causing disruption of the hydrophobic interaction between cytochrome c and cardiolipin, thus releasing cytochrome c [24]. Furthermore, the brain is particularly susceptible to oxidative damage due to its composition of high lipid content. Indeed, ROS play a significant role in the regulation of cell death; however, ROS have recently been reported to induce DNA demethylation via 8-oxoguanine DNA glycosylase-1 (OGG1) [25]. As a result, DNA demethylation induces activation of the reelin gene [26], which has been implicated in enhancing synaptic plasticity by inducing long-term potentiation (LTP) [27], thus indicating that normal levels of ROS may play a role in supporting LTP. Additionally, elimination of ROS negatively impacted neural stem cell proliferation in hippocampal cells indicating that homeostatic levels of ROS may possibly be involved in cell proliferation during growth and development [28]; however additional information is needed in order to elucidate the mechanism behind this.
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\n
\n
2. Mitochondrial dynamics
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Mitochondria were previously thought of as static organelles. Due to advances in molecular biotechnologies, it has been revealed that mitochondria are indeed very dynamic; mitochondria undergo fission and fusion, can vary in morphology, and achieve intracellular movement. Precise execution of these processes is especially vital for proper ATP production, apoptosis, and ROS homeostasis in neurons to properly execute neurotransmission.
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2.1 Fission and fusion
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Fission and fusion are integral processes of cellular homeostasis that maintain proper mitochondrial morphology and turnover. Both are mediated by GTPases in the dynamin family, with rates of occurrence depending on changes in metabolic demands. Undoubtedly, fission is essential for dividing cells in order to maintain an adequate number of mitochondria; however, even in nonproliferating neurons, fission is necessary for cell survival [29]. Dynamin-related protein 1 (Drp1) is the primary GTPase that mediates fission, with its activity controlled by phosphorylation via kinases, primarily on two serine residues. Specifically, phosphorylation at Ser616 promotes fission, while phosphorylation at Ser637 inhibits fission, so balance of Drp1 phosphorylation is crucial for proper fission functionality [30]. Impairment in Drp1 leads to alterations in mitochondrial distribution, with mitochondria accumulation occurring at the soma and reduced density in the dendrites. Conversely, Drp1 overexpression yields an increase in dendritic mitochondria [31]. Hippocampal neurons lacking Drp1 display compromised function of axonal mitochondria due to the inability to maintain ATP levels, recycling at synapses [32]. Prominent regulators of fusion include mitofusion 1 and 2 (Mfn1 and Mfn2, respectively) and optic protein atrophy 1 (Opa1). Mitofusion proteins mediate the outer membrane, while Opa1 regulates the inner membrane; however, both work in coordination in a two-step process to carry out fusion [33, 34].
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Both fission and fusion are enhanced by Bcl-xL, with fission being induced in a Drp1-dependent manner [35]. This is conclusive with a previous study demonstrating the direct interaction of Bcl-xL with Drp1, initiating Drp1-dependent synapse formation in hippocampal cells [36]. When investigated further, this Bcl-xL-Drp1 complex was found to be necessary for presynaptic plasticity by regulating endocytic vesicles [37].
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2.2 Mitochondrial trafficking
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Trafficking, mobility, and docking are intertwining processes that are vital to ensure neurons are equipped with the proper distribution and recycling of mitochondria at axons and synapses throughout the cell’s life span. Figure 1 demonstrates how mitochondria are motile and change morphology in primary hippocampal cells. Mitochondrial trafficking is mediated by intracellular signaling, physiological events, and alterations in metabolic demands. Approximately 70% of mitochondria are stationary, with the remaining 30% motile [38]. Furthermore, five distinct mitochondria motility patterns have been described by Sun’s research group: stationary outside of synapses, docking at synapses, passing though synapses, pausing at synapses for a short amount of time, and pausing for a longer time [39].
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Figure 1.
Mitochondrial movement in primary hippocampal neurons. Primary hippocampal neurons were labeled with mitoRFP, a red fluorescent tag that labels mitochondria. Micrographs were taken at 3 and 4 weeks after seeding. Morphology and location of mitochondria change over time.
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Mechanisms of mitochondrial movement and transport are overall influenced by the polarity of axons, with the positive end directed toward the soma and the negative at the tips. Utilizing this consistent axonal polarity is how microtubule motors drive transport in two directions. Movement away from the soma or anterograde movement is conducted by the ATPase family of kinesins, with kinesin-1 being responsible for mitochondrial transport, specifically in neurons [38, 40]. Kinesin-1 consists of heavy chains (KHC) and light chains, with the heavy chains being the driving force that allows kinesin-1 to function as a motor protein [41]. Retrograde movement or movement toward the soma is driven by dynein. However, it is likely that these movements are coordinated rather than competitive toward each other, as it has been demonstrated that inhibiting kinesin-1 in Drosophila reduces retrograde movement [42].
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Mitochondrial Rho-GTPase, or Miro, is an outer membrane receptor, Ca2+ sensor, and another pertinent regulator of mitochondrial motility due to its ability to anchor kinesin and dynein to the mitochondrial outer membrane [43]. Miro’s anchoring role has been extensively studied in anterograde movement in the motor/adaptor complex formed between KHC and Miro, connected by the protein adaptor milton [44, 45].
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Another important component of mitochondria trafficking is stationary docking. Mitochondrial docking is largely mediated by the axonal outer membrane protein syntaphilin (SNPH) and its interaction with microtubules in the cytoskeleton. This is demonstrated in rodent models in which deletion of the SNPH gene results in an increase in mitochondria motility and reduced density, while overexpression of endogenous or exogenous SNPH abolished mobility [46]. Along with decreasing the percentage of immobile mitochondria, loss of SNPH decreases axonal branching in cortical neurons [47]. This effect is comparable to neurons lacking LKB-1 and NUAK1, which is necessary for axonal specification [48]. The removal of either of these kinases leads to a decrease in the number of stationary mitochondria along with decreased branching. However, overexpression of SNPH in the LKB-1 and NUAK1-null neurons rescued these effects. Collectively, this implies that docking of mitochondria is required for axonal branching and growth [47]. Since Bcl-xL is required for neurite outgrowth [49], it is possible that Bcl-xL exerts this effect by interacting with docking proteins such as syntaphilin. However, the exact role of Bcl-xL in docking mechanisms must be further elucidated.
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3. Alteration of mitochondrial function in brain-associated diseases
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While various brain-associated diseases have different pathophysiologies, there is an underlying similarity that consistently occurs: mitochondrial dysfunction. Throughout these conditions, neurodegeneration is correlated with an energy deficit caused by inefficient operation of the ETC, activation of mitochondria-dependent apoptosis, and accumulation of ROS. In addition, excitotoxicity, which commonly occurs during cerebral ischemia and traumatic brain injury, impairs homeostasis of excitatory neurotransmitter glutamate [50, 51, 52, 53]. Overstimulation of glutamate receptors further leads to Ca2+ release; because mitochondria are one of the key regulators of Ca2+, excessive influx can consequently lead to mitochondrial dysfunction, altered membrane permeabilization, and subsequent cell death [50, 54, 55, 56]. Pathways such as these have been extensively explored in neurological conditions. However, research in the past decade has begun to determine the relationship between brain-associated conditions and mitochondrial dynamics.
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3.1 Parkinson’s disease
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Parkinson’s disease (PD) is a common neurodegenerative disease that has detrimental clinical effects including tremors, impaired gait, and stiffness of limbs [57]. These symptoms are often due to PD’s hallmark characterization of degeneration of the dopaminergic neurons in the substantia nigra. Individuals with PD are vulnerable to increased ROS production due to reduced complex 1 activity, increased lipid oxidation, and altered antioxidant systems [58].
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As several PD-specific proteins impact mitochondrial dynamics, it is possible that the neurodegeneration that occurs with PD is linked to alterations in fission and fusion [59, 60]. Dopaminergic neurons depleted of Drp1 demonstrated decreased mitochondrial mass, impaired motility, and overall neuron loss. Neurons depleted with Drp1 had less mitochondria in the soma and were almost completely depleted from the axons; by not having mitochondria at axons, this can lead to the neurodegeneration due to energy deficits, as synaptic transmission requires a high demand of ATP [61].
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The PINK1/Parkin pathway has been traditionally studied with its roles in mitophagy. Under normal physiological conditions, PINK1 accumulates on the surface of dysfunctional mitochondria to signal Parkin translocation to initiate ubiquitination [62]. However, mutations in PINK1 and Parkin, which have been linked to early onset familial forms of PD, lead to loss of mitochondrial membrane potential, leading to impairment of Parkin’s translocation and thus accumulation of dysfunctional mitochondria [63]. Research in recent years has begun to uncover the role of the PINK1/Parkin pathway in mitochondrial transport. Overexpression of PINK1 phosphorylates Miro, targeting it for ubiquitination and subsequent degradation. This results in the dismantling of the motor/adaptor complex, releasing kinesin and milton from the mitochondrial surface, and leads to halting of mitochondrial motility [64]. It is possible that this system may promote neuroprotection by preventing anterograde transport of mitochondria and allowing PINK1 to accumulate on damaged mitochondria to initiate mitophagy [65]. Furthermore, PINK1 may exert neuroprotection due to its interaction with Bcl-xL [66]. It has been shown that PINK1 phosphorylates Bcl-xL at its Ser62; as a result, this prevents N-terminal cleavage of Bcl-xL or formation of ΔN-Bcl-xL, which has been associated with neuronal death [67, 68, 69]. However, if altered PINK1 expression occurs as a result of genetic mutation, this may lead to dysregulated mitochondrial transport and promotion of apoptosis.
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The presynaptic protein α-synuclein is a major constituent of Lewy bodies, with mutations in its encoding gene, SNCA, being linked to familial PD [70]. Some amount of α-synuclein can localize to the mitochondria, inducing mitochondrial fragmentation, dysfunction, and downregulation of complex 1 activity, potentially contributing to ROS production [71]. Overexpression of α-synuclein results in cytotoxicity due to decreased Bcl-xL expression and increased Bax expression [72]. Shaltouki’s research group recently investigated the role of α-synuclein on mitochondrial dynamics by using multiple PD models. In the postmortem brains of humans with PD, it was observed that protein levels of α-synuclein and Miro were highly upregulated compared to the control brains, while KHC, VDAC, and Mfn2 remained unchanged. Additionally, the β-subunit of ATP synthase was upregulated in the PD brains. When this was further explored in human cell lines and in Drosophila bearing SNCA mutations, neurodegeneration and locomotion defects occurred as a result of the upregulated α-synuclein and subsequent upregulation of Miro. These effects were rescued with a partial reduction of Miro. Interestingly, upregulation of Miro led to delayed mitophagy implying that α-synuclein’s impact on Miro is probably independent of the PINK1/Parkin pathways [73].
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Recently, it was shown that the β-subunit of ATP synthase binds to DJ-1 suggesting that DJ-1 plays a role in increasing ATP efficiency [74]. Mutations in DJ-1 also demonstrate inefficient ATP production, alterations in mitochondrial morphology, and enhanced membrane permeability [74, 75, 76]. Although its functions are not completely understood, DJ-1 has been noted to prevent the aggregation of α-synuclein [77]. As Lewy bodies in PD are primarily a result of α-synuclein aggregation, inhibiting this aggregation may consequently delay Lewy body formation. Thus, therapies targeting DJ-1 may serve as a multi-faceted mechanism for PD treatment.
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3.2 Alzheimer’s disease
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Alzheimer’s disease (AD) is the most common neurodegenerative disease and the leading cause of dementia worldwide [78]. Two hallmark characteristics of AD are the presence of amyloid-beta peptide (Aβ) plaques and tau protein tangles. The formation of Aβ occurs as a consequence of cleavage of amyloid precursor protein (APP), where Aβ peptides can then aggregate into oligomers or fibrils [79]. In the brain, the typical role of tau proteins is to stabilize microtubules; however, in AD, it is suspected that hyperphosphorylation of tau leads to the formation of neurofibril tangles [80]. Both Aβ and tau tangles can cause impairments or interruptions in pathways essential for neuronal survival.
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Reduced transport of axonal mitochondria has been documented in subjects with AD, but there may be multiple mechanisms to cause this disruption [81]. The presence of Aβ reduces bidirectional axonal mitochondrial motility but has a more significant impact on anterograde movement versus retrograde [82, 83]. This may be because Aβ activates GSK-3β, which is a negative regulator of kinesin-1. Phosphorylation of kinesin-1 by GSK-3β can lead to a reduction in mitochondria density [84]. Furthermore, mutations in presenilin-1 (PS1), which is linked to familial AD, promote GSK-3β-mediated kinesin-1 phosphorylation and reductions of anterograde mitochondria transport [84]. Overexpression of tau also has the ability to redirect mitochondrial transport; kinesin-1 encounters tau and is detracted from microtubule tracks, slowing down anterograde movement and increasing the favorability of dynein-mediated retrograde movement [85, 86]. Abnormal fission mechanics have also been observed in AD as a result of alteration in Drp1 function. Neurotoxicity via tau interrupts fission, causing elongated mitochondria and mislocalization of Drp1. This occurs because hyperphosphorylation of tau causes disruptions in actin, preventing actin-based translocation of Drp1 to the mitochondria [87].
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3.3 Stroke
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Cerebral ischemia, or more commonly stroke, is characterized by the decrease or cessation of blood flow to the brain. Consequently, the loss of oxygen and nutrients to neurons causes ATP deficits, apoptosis, and Ca2+ influx. It is not surprising then that mitochondrial dynamics are influenced after ischemic events. This is demonstrated by Zou’s research group as they sought to elucidate how mechanisms of fission and mitophagy are impacted after ischemia [88]. Using a model of middle cerebral artery occlusion (MCAO), Drp1 initially increased but then decreased, implying that ischemia induced fission, but the process was disrupted due to abnormalities in translocation of Drp1 caused by MCAO. Mitophagy is also selectively induced by mild ischemia in a Drp1-dependent manner; this is evident by increased expressions of LC3B and Beclin-1 and decreased p62. Moreover, inhibition of Drp1 led to early onset of apoptotic pathways [88]. This may be supported by transient ischemia models, in which p-Drp1 is upregulated [89, 90]. Interestingly, evidence shows that p-Drp1 at Ser616 may be regulated by PINK1, establishing a link between fission and mitophagy [90]. Like fission, mechanisms of fusion are also impacted by ischemic insult. Mfn2 expression is decreased after MCAO and leads to apoptosis, but when overexpressed, Mfn2 shows an anti-apoptotic effect by modulating Bcl-2 and Bax [91]; these results are conclusive with a similar study, showing that Mfn2 expression is decreased after excitotoxic insult with a subsequent increase in Bax translocation to the mitochondria [92].
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Excitotoxicity via overactivation of glutamate receptors, namely, N-methyl-D-aspartate (NMDA) receptors (NMDARs), is a key player of neuronal death after cerebral ischemia [93]. Thus, uncovering mechanisms effecting NMDARs is an attractive idea for therapeutic agents. While mechanisms of mitochondrial motility are less investigated in models of cerebral ischemia, a recent study has elucidated a novel role of kinesin-1 transport. The heavy chain of kinesin-1 has been shown to bind directly to NMDARs, mediating their transport. By either disassociating this bond or suppressing kinesin-1 expression, this can improve Ca2+ influx and NMDA excitotoxicity resulting from ischemia [94].
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3.4 Traumatic brain injury (TBI)
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Reducing excessive fission that occurs post-TBI is a potential target of neuroprotection to prevent neuronal impairments and death. In rodent models, TBI causes an increase in translocation of Drp1 to the mitochondria, increasing rates of fission. Consequently, this led to neuronal apoptosis, decreased neurogenesis, impaired cognition, and memory defects. When administered with Mdivi-1, a pharmacological inhibitor of Drp1, these negative effects were attenuated, confirming the role of increased Drp1 activity [95, 96]. Interestingly, Pietro’s research group suggest that many molecular responses after severe TBI are opposite from that of mild TBI. Rodents with severe TBI presented with activation of fission as shown by overexpression of Drp1 and FIS1, a protein that binds to Drp1 for anchoring to the mitochondrial outer membrane. Furthermore, expressions of Mfn1 and Mfn2 were downregulated, and there were no changes in Opa1 gene and protein expressions, demonstrating an inhibition of fusion as a result of severe TBI. Additionally, the increase of dysfunctional mitochondria led to a subsequent overexpression of PINK1 and PARK2, triggering mitophagy. Conversely, mild TBI demonstrated activation of fusion with inhibition of fission; together, this did not change PINK1 or PARK2 gene expressions, thus showing no difference in mitophagy [97].
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4. The effect of lifestyle factors on mitochondria function and dynamics
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4.1 Exercise and mitochondria
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Traditionally observed at a large, systemic level, research in recent years has begun to investigate the effect of exercise on mitochondrial function and dynamic processes. Furthermore, encouraging results have been observed by analyzing these effects in physiological processes in the brain and pathological conditions such as Alzheimer’s disease and Parkinson’s disease [98, 99, 100, 101]. After old and young mice were subjected to 6 weeks of treadmill exercise, old mice were found to respond positively, showing attenuated activity of coupled complexes I–III of the ETC [99].
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Exercise regulates mitochondrial fission and fusion proteins such as Opa1, Mfn2, and Drp1 and enhances mitochondrial biogenesis via upregulation of mitochondrial DNA. This suggests a role for exercise in maintaining a healthy population of mitochondria [98, 101]. In addition, rodents that undergo physical exercise demonstrate improved cognitive and exploratory behaviors along with improved mitochondrial redox homeostasis and mitochondria-mediated brain energy metabolism [102, 103, 104].
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Exercise has been shown to regulate apoptosis through the regulation of the Bcl-2 family in various models and tissues. As previously discussed, age can have a significant role in mitochondrial dysfunction. A study conducted by Kim et al. investigated how hippocampal neurogenesis and apoptosis were affected by treadmill exercise in young and old-aged rats [105]. Expressions of caspase-3, Bax, Bid, and Bcl-2 were all increased in old mice. After being subjected to treadmill exercise for 30 minutes, once per day for 6 weeks, old rats exhibited further enhancement of Bcl-2 expression, along with decreased expressions of caspase-3, Bax, and Bid. Interestingly, exercise did not impact expressions of Bcl-2, Bax, or caspase-3 in young mice. These results implicate that aerobic exercise may be especially important during aging to exert neuroprotective properties. Aboutaleb et al. examined the effect exercise had on the ratio of Bax/Bcl-2 proteins in hippocampal CA3 cells after ischemia. Ischemic insult led to an increase in caspase-3 and decrease in Bcl-2, thus an increase in Bax/Bcl-2 ratio. However, rats pre-subjected to exercise showed reduced levels of caspase-3 and attenuation of Bax/Bcl-2 ratio [106]. Similar results have been seen in a rodent TBI model, in which treadmill exercise lowered the Bax/Bcl-2 ratio and decreased the levels of active caspase-3 [107]. Likewise, endurance exercise exerted neuroprotection in PD models by modulation of Mcl-1, Bcl-2, and apoptosis-inducing factor [108]. Although Bcl-2 has primarily been investigated in rodent exercise models, exercise results in a negative regulatory effect on caspase-3 [105, 106, 107] which may reduce post-translational cleavage of Bcl-xL supporting neuronal survival [67, 69].
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Since the Bcl-2 family proteins are present in the mitochondria throughout all tissue types, it is plausible that protective effects observed in non-neuronal mitochondria are indeed simultaneously occurring in neuronal mitochondria. For example, due to the correlation between diabetes and cardiovascular disease, Cheng et al. examined the relationship between apoptosis, cardiomyocytes, and aerobic exercise in streptozotocin (STZ)-induced diabetic rats. The STZ rats had significantly lower amounts of Bcl-2, Bcl-xL, and p-Bad and higher levels of caspase-3 than controls. However, these levels were all rescued when subjected to aerobic exercise [109]. These results implicate the ability of aerobic exercise to regulate apoptosis and exert cellular protection, even during chronic conditions.
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4.2 Non-neurological conditions and mitochondrial consequences
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As chronic diseases such as diabetes and obesity become more prevalent worldwide, research is uncovering the relationship between traditional non-neurological and brain-associated diseases. However, many interventions in studies concerning chronic diseases observe outcomes on a macrolevel and may not consider molecular effects. For these reasons, it is important to uncover how these non-neurological chronic conditions are impacting the mitochondria. By elucidating these mechanisms, this can serve as an additional, albeit perhaps overlooked, means of prevention against brain pathology.
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Besides disrupting the uptake of glucose, diabetes can also damage the function, population, and morphology of mitochondria in neurons, potentially contributing to impaired cognition later in life [110, 111, 112, 113]. Metabolic pathways that can lead to energy failure in mitochondria as well as prevent antioxidant interception are affected by diabetes. In the diabetic brain, mitochondrial perturbation can result in a lack of neuronal energy that will alter synaptic function and eventually cause the neurons to degenerate [114]. The effect of energy impairments is demonstrated in a cross-sectional study in which adolescents with type 2 diabetes had slower conversion rates of ADP to ATP than obese and lean controls. The explanation for this effect was suspected to be due to decreased blood flow, thus causing alterations in oxygen delivery [115]. Studies have shown that diabetes can modify fission mechanisms in rodent models. Although amounts of Mfn1 and Opa1 remain unchanged during diabetes, Drp1 mRNA is increased. Furthermore, there is an increase of translocation of Drp1 to the mitochondria in diabetes [110, 113]. This increase in translocation is due to GSK3β-mediated phosphorylation at Ser616 of Drp1. The combination of unchanged Mfn1 and Opa1 with increased Drp1 proposes that the disproportion between fission and fusion proteins contributes to mitochondrial dysfunction in rats with diabetes. This was further evident by altered mitochondrial morphology and density. Elevated levels of Drp1 can lead to mitochondrial fragmentation that is conducive to damage in the synapses of neurons, contributing to impairments in learning and memory [116]. Beyond alterations in fission and fusion, decreased ATP production and activity of complex I were observed in the diabetic hippocampus. Moreover, glutathione and ascorbate levels were decreased, suggesting that diabetes impairs mitochondrial antioxidant systems [110]. These results are supported by a study that found decreased coenzyme Q9 and ATPase activity in the mitochondria of diabetic rats [117].
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Obesity is known to increase the risk of developing diabetes, cardiovascular disease, and neurological conditions. Indeed, obese animal models are often utilized to study insulin resistance. A characteristic of obesity is chronic inflammation and oxidative stress, so it is not surprising that mitochondrial dysfunction occurs throughout the body, including the brain [114]. Specifically, brain mitochondria of obese rats induced via a high-fat diet have repeatedly demonstrated a shift to pro-apoptotic pathways, as shown by elevated Bax expressions, lowered Bcl-2, and a higher Bax/Bcl-2 ratio [118, 119]. The detrimental effects of obesity continue to be demonstrated in the brain by upregulating production of ROS and alteration of mitochondrial membrane potential [120, 121, 122].
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4.3 Diet and mitochondria
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Diet, including intake of specific nutrients and overall encompassing dietary patterns, is a driver of maintaining cellular processes throughout the body. Treatment of diseases via diet is appealing due to the ability of nutrients to cross the blood brain barriers and ease of accessibility. Specifically, it is important to consider how an individual’s overall dietary pattern impacts cellular processes. Dietary patterns, including composition of macronutrients and caloric provision, have been studied regarding efficacy in neuroprotection [123, 124, 125].
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Caloric restriction has been implicated in the protection against several pathological brain conditions in various animal models such as AD and PD and under conditions of excitotoxicity [126, 127, 128, 129, 130]. Caloric restriction has been shown to confer protection from neurodegeneration by improving mitochondrial redox status by reducing ROS production localized to complex I of the ETC [131]. Recent evidence suggests that caloric restriction may prevent formation of ROS via upregulation of antioxidants such as mitochondrial superoxide dismutase 2 (SOD2) and glutathione [132]. Caloric restriction has also been reported to upregulate antioxidants localized to the plasma membrane such as coenzyme Q10 and α-tocopherol via an increase in redox enzymes that are capable of reducing these molecules back to their antioxidant form [133]. Due to coenzyme Q10’s pivotal role as an electron carrier in the ETC, we speculate that caloric restriction may be beneficial to maintain redox balance in the mitochondrial membrane. Additionally, mRNA expressions of Bcl-2 and Bcl-xL were also reported to be upregulated in the ipsilateral cortex region of mice placed on caloric restriction against TBI [134], indicating that caloric restriction may prevent TBI-induced neuronal loss. Furthermore, caloric restriction improves mitochondrial function by enhancing ATP levels in aging mice [135]. Mice placed on caloric restriction for 6 months had increased mitochondrial biogenesis and increased levels of cytochrome c oxidase and citrate synthase activity, enhancing mitochondrial respiration [136]. Caloric restriction may enhance mitochondrial metabolism by also upregulating the activity of complexes I, III, and IV [128]. Interestingly, recent evidence shows that caloric restriction enhances expression of brain-derived neurotrophic factor (BDNF) [137, 138], which has been reported to regulate mitochondrial mobility and enhance presynaptic docking [139]. However, the mechanisms of how caloric restriction mediates BDNF expression are still unclear. Clinical trials in which older adults are placed on caloric restriction consistently yield positive results, such as improved memory and enhanced gray matter [140, 141]. Additionally, caloric restriction attenuated behavioral dysfunction in a model of PD in adult rhesus monkeys [130]. Taken together, these studies point toward caloric restriction mediating biological markers of chronic disease such as oxidative stress and supporting mitochondrial function by enhancing ATP metabolism and possibly lessening clinical symptoms associated with neurodegeneration.
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The ketogenic diet, popular for its high-fat and very low carbohydrate pattern, has recently been implicated in protection of the brain through apoptotic pathways. Various mammalian animals placed on the ketogenic diet show decreased rates of apoptotic stimuli in neuronal cells via downregulation of mitochondrial cytochrome c release and active caspase-3 both in seizures [142, 143] and TBI models [144], respectively. The decrease in translocation of cytochrome c from the mitochondria to the cytosol may be through the regulation of Bcl-2. One study found that both a high carbohydrate and a high ketogenic diet upregulate Bcl-2 in cortical neurons after focal cerebral ischemia; however, the ketogenic diet displayed higher upregulation [145], indicating that the ketogenic diet may be more efficient in regulating apoptosis than a high carbohydrate diet. The ketogenic diet may play an additional role in cell death and survival pathways, as it has been noted to protect hippocampal cells from death by preventing the interaction between Bad and Bcl-xL [146]. The ketogenic diet further supports neuronal energy metabolism by maintaining mitochondrial morphology, enhancing biogenesis of mitochondria, and improving mitochondrial respiration [147, 148, 149, 150, 151]. After neurotoxic insult, the ketogenic diet enhanced complex I-driven oxygen consumption and prevented loss of complex II–III function, implicating the ketogenic diet’s ability to improve the activity of the ETC [147, 149]. Likewise, the ketogenic diet attenuates mitochondrial oxidative stress levels in both in vitro and in vivo model, which prevents energy deficit associated with brain cell damage [147, 149, 151]. Interestingly, the ketogenic diet has also been shown to upregulate Beclin-1 [142] and Drp1 [148], suggesting that the ketogenic diet may be able to control mitochondrial population by regulating autophagy and mitochondrial fission, respectively.
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5. Conclusions
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Mitochondria are well established in their role with ATP production, apoptosis, ROS homeostasis, and intracellular ion signaling. Research in recent years has recognized that proper execution of these processes is reliant on the mitochondria’s dynamic capabilities. In this chapter we have discussed mechanisms of mitochondrial morphology, degradation, and trafficking, as well as the relationship between these processes and pathological brain conditions. Utilizing lifestyle factors, such as exercise and diet, can serve as a neuroprotective strategy by targeting neuronal mitochondrial dynamics. Implementing lifestyle changes serves as an accessible treatment that is easily translated from bench to bedside.
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Acknowledgments
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This research was funded by Summer Research Support by the College of Human Environmental Sciences, the University of Alabama.
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Conflict of interest
The authors declare no conflict of interest.
\n',keywords:"mitochondria, neuroprotection, Bcl-2, exercise, diet",chapterPDFUrl:"https://cdn.intechopen.com/pdfs/69122.pdf",chapterXML:"https://mts.intechopen.com/source/xml/69122.xml",downloadPdfUrl:"/chapter/pdf-download/69122",previewPdfUrl:"/chapter/pdf-preview/69122",totalDownloads:328,totalViews:0,totalCrossrefCites:0,dateSubmitted:"May 6th 2019",dateReviewed:"August 29th 2019",datePrePublished:"September 19th 2019",datePublished:"November 26th 2020",dateFinished:null,readingETA:"0",abstract:"The brain requires vast amounts of energy to carry out neurotransmission; indeed, it is responsible for approximately one-fifth of the body’s energy consumption. Therefore, in order to understand functions of brain cells under both normal and pathological conditions, it is critical to elucidate dynamics of intracellular energy. The mitochondrion is the key intercellular organelle that controls neuronal energy and survival. Numerous studies have reported a correlation between altered mitochondrial function and brain-associated diseases; thus mitochondria may serve as a promising target for treating these conditions. In this chapter, we will discuss the mechanisms of mitochondrial production, movement, and degradation in order to understand accessibility of energy during physiological and pathological conditions of the brain. While research targeting molecular dynamics is promising, translation into clinical relevance based on bench research is challenging. For these reasons, we will also summarize lifestyle factors, including interventions and chronic comorbidities that disrupt mitochondrial dynamics. By determining lifestyle factors that are readily accessible, we can propose a new viewpoint for a synergistic and translational approach for neuroprotection.",reviewType:"peer-reviewed",bibtexUrl:"/chapter/bibtex/69122",risUrl:"/chapter/ris/69122",signatures:"Katheryn Broman, Abigail U. Davis, Jordan May and Han-A Park",book:{id:"8087",title:"Neuroprotection",subtitle:"New Approaches and Prospects",fullTitle:"Neuroprotection - New Approaches and Prospects",slug:"neuroprotection-new-approaches-and-prospects",publishedDate:"November 26th 2020",bookSignature:"Matilde Otero-Losada, Francisco Capani and Santiago Perez Lloret",coverURL:"https://cdn.intechopen.com/books/images_new/8087.jpg",licenceType:"CC BY 3.0",editedByType:"Edited by",editors:[{id:"193560",title:"Dr.",name:"Matilde",middleName:null,surname:"Otero-Losada",slug:"matilde-otero-losada",fullName:"Matilde Otero-Losada"}],productType:{id:"1",title:"Edited Volume",chapterContentType:"chapter",authoredCaption:"Edited by"}},authors:[{id:"304239",title:"Dr.",name:"Han-A",middleName:null,surname:"Park",fullName:"Han-A Park",slug:"han-a-park",email:"hpark36@ches.ua.edu",position:null,institution:null},{id:"309383",title:"MSc.",name:"Katheryn",middleName:null,surname:"Broman",fullName:"Katheryn Broman",slug:"katheryn-broman",email:"kabroman@crimson.ua.edu",position:null,institution:{name:"University of Alabama, Tuscaloosa",institutionURL:null,country:{name:"United States of America"}}},{id:"309385",title:"BSc.",name:"Abigail",middleName:null,surname:"Davis",fullName:"Abigail Davis",slug:"abigail-davis",email:"audavis@crimson.ua.edu",position:null,institution:{name:"University of Alabama, Tuscaloosa",institutionURL:null,country:{name:"United States of America"}}},{id:"309386",title:"Mr.",name:"Jordan",middleName:null,surname:"May",fullName:"Jordan May",slug:"jordan-may",email:"jmmay4@crimson.ua.edu",position:null,institution:{name:"University of Alabama, Tuscaloosa",institutionURL:null,country:{name:"United States of America"}}}],sections:[{id:"sec_1",title:"1. Introduction",level:"1"},{id:"sec_2",title:"2. Mitochondrial dynamics",level:"1"},{id:"sec_2_2",title:"2.1 Fission and fusion",level:"2"},{id:"sec_3_2",title:"2.2 Mitochondrial trafficking",level:"2"},{id:"sec_5",title:"3. Alteration of mitochondrial function in brain-associated diseases",level:"1"},{id:"sec_5_2",title:"3.1 Parkinson’s disease",level:"2"},{id:"sec_6_2",title:"3.2 Alzheimer’s disease",level:"2"},{id:"sec_7_2",title:"3.3 Stroke",level:"2"},{id:"sec_8_2",title:"3.4 Traumatic brain injury (TBI)",level:"2"},{id:"sec_10",title:"4. The effect of lifestyle factors on mitochondria function and dynamics",level:"1"},{id:"sec_10_2",title:"4.1 Exercise and mitochondria",level:"2"},{id:"sec_11_2",title:"4.2 Non-neurological conditions and mitochondrial consequences",level:"2"},{id:"sec_12_2",title:"4.3 Diet and mitochondria",level:"2"},{id:"sec_14",title:"5. Conclusions",level:"1"},{id:"sec_15",title:"Acknowledgments",level:"1"},{id:"sec_18",title:"Conflict of interest",level:"1"}],chapterReferences:[{id:"B1",body:'\nSmith GM, Gallo G. The role of mitochondria in axon development and regeneration. Developmental Neurobiology. 2018;78(3):221-237\n'},{id:"B2",body:'\nHarris JJ, Jolivet R, Attwell D. Synaptic energy use and supply. Neuron. 2012;75(5):762-777\n'},{id:"B3",body:'\nSkou JC. The identification of the sodium pump. Bioscience Reports. 2004;24(4-5):436-451\n'},{id:"B4",body:'\nPivovarov AS, Calahorro F, Walker RJ. Na(+)/K(+)-pump and neurotransmitter membrane receptors. Invertebrate Neuroscience. 2018;19(1):1\n'},{id:"B5",body:'\nPathak D, Shields LY, Mendelsohn BA, Haddad D, Lin W, et al. The role of mitochondrially derived ATP in synaptic vesicle recycling. The Journal of Biological Chemistry. 2015;290(37):22325-22336\n'},{id:"B6",body:'\nKroemer G, Dallaporta B, Resche-Rigon M. 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Effect of streptozotocin-induced diabetes on rat brain mitochondria. Journal of Neuroendocrinology. 2004;16(1):32-38\n'},{id:"B113",body:'\nHuang S, Wang Y, Gan X, Fang D, Zhong C, et al. Drp1-mediated mitochondrial abnormalities link to synaptic injury in diabetes model. Diabetes. 2015;64(5):1728-1742\n'},{id:"B114",body:'\nSripetchwandee J, Chattipakorn N, Chattipakorn SC. Links between obesity-induced brain insulin resistance, brain mitochondrial dysfunction, and dementia. Frontiers in Endocrinology (Lausanne). 2018;9:496\n'},{id:"B115",body:'\nCree-Green M, Gupta A, Coe GV, Baumgartner AD, Pyle L, et al. Insulin resistance in type 2 diabetes youth relates to serum free fatty acids and muscle mitochondrial dysfunction. Journal of Diabetes and Its Complications. 2017;31(1):141-148\n'},{id:"B116",body:'\nVantaggiato C, Castelli M, Giovarelli M, Orso G, Bassi MT, et al. The fine tuning of Drp1-dependent mitochondrial remodeling and autophagy controls neuronal differentiation. Frontiers in Cellular Neuroscience. 2019;13:120\n'},{id:"B117",body:'\nMoreira PI, Santos MS, Sena C, Seica R, Oliveira CR. Insulin protects against amyloid beta-peptide toxicity in brain mitochondria of diabetic rats. Neurobiology of Disease. 2005;18(3):628-637\n'},{id:"B118",body:'\nSa-Nguanmoo P, Tanajak P, Kerdphoo S, Jaiwongkam T, Pratchayasakul W, et al. SGLT2-inhibitor and DPP-4 inhibitor improve brain function via attenuating mitochondrial dysfunction, insulin resistance, inflammation, and apoptosis in HFD-induced obese rats. Toxicology and Applied Pharmacology. 2017;333:43-50\n'},{id:"B119",body:'\nSa-Nguanmoo P, Tanajak P, Kerdphoo S, Satjaritanun P, Wang X, et al. FGF21 improves cognition by restored synaptic plasticity, dendritic spine density, brain mitochondrial function and cell apoptosis in obese-insulin resistant male rats. Hormones and Behavior. 2016;85:86-95\n'},{id:"B120",body:'\nPintana H, Sripetchwandee J, Supakul L, Apaijai N, Chattipakorn N, et al. Garlic extract attenuates brain mitochondrial dysfunction and cognitive deficit in obese-insulin resistant rats. Applied Physiology, Nutrition, and Metabolism. 2014;39(12):1373-1379\n'},{id:"B121",body:'\nPratchayasakul W, Sa-Nguanmoo P, Sivasinprasasn S, Pintana H, Tawinvisan R, et al. Obesity accelerates cognitive decline by aggravating mitochondrial dysfunction, insulin resistance and synaptic dysfunction under estrogen-deprived conditions. Hormones and Behavior. 2015;72:68-77\n'},{id:"B122",body:'\nWang D, Yan J, Chen J, Wu W, Zhu X, et al. Naringin improves neuronal insulin signaling, brain mitochondrial function, and cognitive function in high-fat diet-induced obese mice. Cellular and Molecular Neurobiology. 2015;35(7):1061-1071\n'},{id:"B123",body:'\nWlodarek D. Role of ketogenic diets in neurodegenerative diseases (Alzheimer’s disease and Parkinson’s disease). Nutrients. 2019;11(1):169\n'},{id:"B124",body:'\nAgarwal P, Wang Y, Buchman AS, Holland TM, Bennett DA, et al. MIND diet associated with reduced incidence and delayed progression of Parkinsonism in old age. The Journal of Nutrition, Health and Aging. 2018;22(10):1211-1215\n'},{id:"B125",body:'\nMorris MC, Tangney CC, Wang Y, Sacks FM, Barnes LL, et al. MIND diet slows cognitive decline with aging. Alzheimer’s and Dementia. 2015;11(9):1015-1022\n'},{id:"B126",body:'\nDuan W, Mattson MP. Dietary restriction and 2-deoxyglucose administration improve behavioral outcome and reduce degeneration of dopaminergic neurons in models of Parkinson’s disease. Journal of Neuroscience Research. 1999;57(2):195-206\n'},{id:"B127",body:'\nAnson RM, Guo Z, de Cabo R, Iyun T, Rios M, et al. Intermittent fasting dissociates beneficial effects of dietary restriction on glucose metabolism and neuronal resistance to injury from calorie intake. Proceedings of the National Academy of Sciences of the United States of America. 2003;100(10):6216-6220\n'},{id:"B128",body:'\nAmigo I, Menezes-Filho SL, Luevano-Martinez LA, Chausse B, Kowaltowski AJ. Caloric restriction increases brain mitochondrial calcium retention capacity and protects against excitotoxicity. Aging Cell. 2017;16(1):73-81\n'},{id:"B129",body:'\nHalagappa VK, Guo Z, Pearson M, Matsuoka Y, Cutler RG, et al. Intermittent fasting and caloric restriction ameliorate age-related behavioral deficits in the triple-transgenic mouse model of Alzheimer’s disease. Neurobiology of Disease. 2007;26(1):212-220\n'},{id:"B130",body:'\nMaswood N, Young J, Tilmont E, Zhang Z, Gash DM, et al. Caloric restriction increases neurotrophic factor levels and attenuates neurochemical and behavioral deficits in a primate model of Parkinson’s disease. Proceedings of the National Academy of Sciences of the United States of America. 2004;101(52):18171-18176\n'},{id:"B131",body:'\nSanz A, Caro P, Ibanez J, Gomez J, Gredilla R, et al. Dietary restriction at old age lowers mitochondrial oxygen radical production and leak at complex I and oxidative DNA damage in rat brain. Journal of Bioenergetics and Biomembranes. 2005;37(2):83-90\n'},{id:"B132",body:'\nHu Y, Zhang M, Chen Y, Yang Y, Zhang JJ. Postoperative intermittent fasting prevents hippocampal oxidative stress and memory deficits in a rat model of chronic cerebral hypoperfusion. European Journal of Nutrition. 2019;58(1):423-432\n'},{id:"B133",body:'\nHyun DH, Emerson SS, Jo DG, Mattson MP, de Cabo R. Calorie restriction up-regulates the plasma membrane redox system in brain cells and suppresses oxidative stress during aging. Proceedings of the National Academy of Sciences of the United States of America. 2006;103(52):19908-19912\n'},{id:"B134",body:'\nLoncarevic-Vasiljkovic N, Milanovic D, Pesic V, Tesic V, Brkic M, et al. Dietary restriction suppresses apoptotic cell death, promotes Bcl-2 and Bcl-xl mRNA expression and increases the Bcl-2/Bax protein ratio in the rat cortex after cortical injury. Neurochemistry International. 2016;96:69-76\n'},{id:"B135",body:'\nLin AL, Coman D, Jiang L, Rothman DL, Hyder F. Caloric restriction impedes age-related decline of mitochondrial function and neuronal activity. Journal of Cerebral Blood Flow and Metabolism. 2014;34(9):1440-1443\n'},{id:"B136",body:'\nCerqueira FM, Cunha FM, Laurindo FR, Kowaltowski AJ. Calorie restriction increases cerebral mitochondrial respiratory capacity in a NO*-mediated mechanism: Impact on neuronal survival. Free Radical Biology and Medicine. 2012;52(7):1236-1241\n'},{id:"B137",body:'\nDuan W, Lee J, Guo Z, Mattson MP. 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Annals of Clinical and Laboratory Science. 2009;39(1):76-83\n'},{id:"B145",body:'\nPuchowicz MA, Zechel JL, Valerio J, Emancipator DS, Xu K, et al. Neuroprotection in diet-induced ketotic rat brain after focal ischemia. Journal of Cerebral Blood Flow and Metabolism. 2008;28(12):1907-1916\n'},{id:"B146",body:'\nNoh HS, Kim YS, Kim YH, Han JY, Park CH, et al. Ketogenic diet protects the hippocampus from kainic acid toxicity by inhibiting the dissociation of bad from 14-3-3. Journal of Neuroscience Research. 2006;84(8):1829-1836\n'},{id:"B147",body:'\nGreco T, Glenn TC, Hovda DA, Prins ML. Ketogenic diet decreases oxidative stress and improves mitochondrial respiratory complex activity. Journal of Cerebral Blood Flow and Metabolism. 2016;36(9):1603-1613\n'},{id:"B148",body:'\nHasan-Olive MM, Lauritzen KH, Ali M, Rasmussen LJ, Storm-Mathisen J, et al. A ketogenic diet improves mitochondrial biogenesis and bioenergetics via the PGC1alpha-SIRT3-UCP2 axis. Neurochemical Research. 2019;44(1):22-37\n'},{id:"B149",body:'\nMaalouf M, Sullivan PG, Davis L, Kim DY, Rho JM. Ketones inhibit mitochondrial production of reactive oxygen species production following glutamate excitotoxicity by increasing NADH oxidation. Neuroscience. 2007;145(1):256-264\n'},{id:"B150",body:'\nNylen K, Velazquez JL, Sayed V, Gibson KM, Burnham WM, et al. The effects of a ketogenic diet on ATP concentrations and the number of hippocampal mitochondria in Aldh5a1(−/−) mice. Biochimica et Biophysica Acta. 2009;1790(3):208-212\n'},{id:"B151",body:'\nYao J, Chen S, Mao Z, Cadenas E, Brinton RD. 2-Deoxy-D-glucose treatment induces ketogenesis, sustains mitochondrial function, and reduces pathology in female mouse model of Alzheimer’s disease. PLoS One. 2011;6(7):e21788\n'}],footnotes:[],contributors:[{corresp:null,contributorFullName:"Katheryn Broman",address:null,affiliation:'
Department of Human Nutrition and Hospitality Management, College of Human Environmental Sciences, The University of Alabama, Tuscaloosa, AL, USA
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Department of Human Nutrition and Hospitality Management, College of Human Environmental Sciences, The University of Alabama, Tuscaloosa, AL, USA
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