Section 2 Energy Metabolism

**9**

**Chapter 2**

*Yuan Lu*

**Abstract**

humans.

glucose, metabolic disorders

**1. Introduction**

Metabolism

Brown Adipose Tissue Energy

The brown adipose tissue (BAT) evolved as a specialized thermogenic organ in mammals. Nutrients (i.e., fatty acids and glucose) from the intracellular storage and peripheral tissues are critical to the BAT thermogenic function. The BAT converts the chemical energy stored in nutrients to thermo energy through UCP1-mediated nonshivering thermogenesis (NST). Activated BAT contributes significantly to the whole body energy substrate homeostasis. It is now well-recognized that adult humans possess BAT with functional thermoactivity. Thus, BAT energy metabolism has a significant therapeutic potential in the management of metabolic disorders, such as obesity, insulin resistance, type 2 diabetes, and lipid abnormality in

**Keywords:** brown adipose tissue, brown adipocyte, metabolism, fatty acid,

disorders, such as diabetes and obesity in humans [8–14, 16, 17, 21, 27].

Brown adipocyte, which is endowed with mitochondria, is the most important thermogenic functional unit of BAT [4, 28]. In the mitochondrion of brown

Brown adipose tissue (BAT) evolved as a specialized thermogenic organ in the modern eutherian mammals, including *Homo sapiens* [1–7]. The main function of BAT is mediating adaptive thermogenesis or nonshivering thermogenesis (NST), when mammals are challenged in the cold environment. The NST function is a critical adaptation, which helps to maintain the homeothermy in mammals and gives them the evolutionary advantage to survive in the cold habitat, and to thrive from the Artic to the Antarctic region [3, 4]. During the cold challenge, BAT metabolizes nutrients (i.e., fatty acids and glucose) and converts the chemical energy to thermo energy through NST. In addition to its thermogenic function, activated BAT also contributes significantly to whole body energy substrate homeostasis. When activated, BAT presents physiologically significant benefit in fatty acids and glucose homeostasis as well as insulin sensitivity in mammals [8–24]. It is now wellrecognized that adult humans possess BAT, which has functional thermoactivity [8–10, 25]. Cold challenge-activated BAT is detected mainly in the supraclavicular, paravertebral, and cervical regions in adult humans [9, 10, 25, 26]. Accumulating evidences indicate that BAT function is inversely associated with age, body mass index (BMI), and diabetic status in adult humans, which indicates that the activation of BAT has potential translational implication in the management of metabolic

#### **Chapter 2**

## Brown Adipose Tissue Energy Metabolism

*Yuan Lu*

#### **Abstract**

The brown adipose tissue (BAT) evolved as a specialized thermogenic organ in mammals. Nutrients (i.e., fatty acids and glucose) from the intracellular storage and peripheral tissues are critical to the BAT thermogenic function. The BAT converts the chemical energy stored in nutrients to thermo energy through UCP1-mediated nonshivering thermogenesis (NST). Activated BAT contributes significantly to the whole body energy substrate homeostasis. It is now well-recognized that adult humans possess BAT with functional thermoactivity. Thus, BAT energy metabolism has a significant therapeutic potential in the management of metabolic disorders, such as obesity, insulin resistance, type 2 diabetes, and lipid abnormality in humans.

**Keywords:** brown adipose tissue, brown adipocyte, metabolism, fatty acid, glucose, metabolic disorders

#### **1. Introduction**

Brown adipose tissue (BAT) evolved as a specialized thermogenic organ in the modern eutherian mammals, including *Homo sapiens* [1–7]. The main function of BAT is mediating adaptive thermogenesis or nonshivering thermogenesis (NST), when mammals are challenged in the cold environment. The NST function is a critical adaptation, which helps to maintain the homeothermy in mammals and gives them the evolutionary advantage to survive in the cold habitat, and to thrive from the Artic to the Antarctic region [3, 4]. During the cold challenge, BAT metabolizes nutrients (i.e., fatty acids and glucose) and converts the chemical energy to thermo energy through NST. In addition to its thermogenic function, activated BAT also contributes significantly to whole body energy substrate homeostasis. When activated, BAT presents physiologically significant benefit in fatty acids and glucose homeostasis as well as insulin sensitivity in mammals [8–24]. It is now wellrecognized that adult humans possess BAT, which has functional thermoactivity [8–10, 25]. Cold challenge-activated BAT is detected mainly in the supraclavicular, paravertebral, and cervical regions in adult humans [9, 10, 25, 26]. Accumulating evidences indicate that BAT function is inversely associated with age, body mass index (BMI), and diabetic status in adult humans, which indicates that the activation of BAT has potential translational implication in the management of metabolic disorders, such as diabetes and obesity in humans [8–14, 16, 17, 21, 27].

Brown adipocyte, which is endowed with mitochondria, is the most important thermogenic functional unit of BAT [4, 28]. In the mitochondrion of brown adipocyte, energy generated from nutrients is initially stored as proton gradient membrane potential across the mitochondrial inner membrane, and then converted directly to thermo energy by the uncoupling protein 1 (UCP1)-mediated proton flow [4, 29]. In BAT, brown adipocytes are surrounded by abundant capillaries. The heat generated in the brown adipocyte mitochondria can be quickly distributed by the blood flow to maintain the steady core body temperature in mammals [4]. In addition to classical brown adipocytes, beige/brite adipocytes can be induced from the white adipocytes to conduct thermogenesis upon the cold challenge or catecholamine stimulation [26, 30, 31]. This process is termed as "browning" [26, 30, 31]. Thermogenesis in beige/brite adipocytes can also contribute to mammals' body temperature maintenance and whole body metabolism [26, 30, 31]. Beige/brite adipose tissue generates heat through both UCP1-independent thermogenesis, including calcium cycling in and out of endoplasmic reticulum, futile cycle between creatine and phosphocreatine, and UCP1-dependent thermogenesis in mitochondria [26, 30–33]. The energy metabolism is essential for the optimal UCP1-mediated mitochondrial thermogenic function of brown and beige/ brite adipocytes [4, 34, 35]. In this chapter, we discuss the importance of energy metabolism in maintaining brown adipocyte thermogenic function and the proceeding of targeting metabolic disorder through BAT activation in human studies.

#### **2. Fatty acid metabolism in brown adipocytes**

Thermogenic brown adipocyte possesses a high capacity for fatty acid β-oxidation that has been reported in both rodent and humans [4, 12, 14, 36]. Fatty acids serve as the activator for UCP1, which is a fatty acid/H+ symporter directly mediating proton flow and thermogenesis [37]. Fatty acids also serve as the main energy substrate mediating the uncoupling and thermogenic function in brown adipocytes [2, 36–41]. In addition, fatty acids can enhance brown adipocyte thermogenic capacity through the nuclear receptor peroxisome proliferator-activated receptors (PPARs), which are the master transcription regulators for the expression of genes involved in lipid metabolism, oxidative phosphorylation, and the key thermogenic protein UCP1 in brown adipocytes [42, 43].

Intracellular fatty acids are stored in the format of triglyceride in the heterogeneous multilocular lipid droplets in the brown adipocyte [4]. Upon the cold challenge, triglycerides stored in the brown adipocyte lipid droplet are lipolysed. Triglyceride lipolysis is a sequential process that involves different enzymes, resulting in the liberation of glycerol and fatty acids for heat production [44]. The lipid droplet is composed of triglycerides and cholesterol esters, which are surrounded by a monolayer of phospholipids [44]. Important proteins with regulatory and enzymatic functions, including perilipin and CGI58, coexist on the phospholipid monolayer to regulate the lipid trafficking and other functions of the lipid droplet [44–47]. Perilipin stabilizes the lipid droplet and prevents it from lipolysis under basal condition. Upon cold challenge, β-adrenergic stimulation leads to the activation of G-protein-coupled receptor (GPCR) and adenylate cyclase, which subsequently increases the cAMP level in brown adipocyte [48]. cAMP then activates protein kinase A (PKA), which phosphorylates perilipin at its serine residues [49–52]. The phosphorylated perilipin releases CGI-58, an adipose triglyceride lipase (ATGL)-activating protein. CGI-58, subsequently, binds and actives ATGL. Activated ATGL hydrolyzes triglycerides and generates free fatty acids and diglycerides [50, 53–55]. Upon the cold challenge, PKA also phosphorylates serine residues on another key lipolysis enzyme hormone sensitive lipase (HSL) [56]. Although HSL is capable of hydrolyzing triglycerides, diglycerides, monoglycerides,

**11**

*Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

free fatty acids and glycerol [57].

of age with an average BMI of 24.5 kg/m2

the BAT [58].

cytes upon activation.

and a broad array of other lipid substrate, it is the rate-limiting enzyme for hydrolyzing the diglycerides *in vivo* [56]. The phosphorylated HSL catalyzes the diglycerides and generates free fatty acids and monoglycerides [56]. As the last step of lipolysis, monoglycerides is hydrolyzed by monoacylglycerol lipase (MGL) to form

Given the importance of fatty acid in brown adipocyte thermogenic function, it is reasonable to predict that the deficiency of the key lipolysis enzymes and fatty acid transportation proteins in brown adipocytes can lead to a defected BAT thermogenic function. Human and rodent *in vivo* studies using both pharmacological and genetic approaches to manipulate triglyceride lipolysis processes were reported [36, 39, 58–60]. In the initial studies, nicotinic acid (NiAc) was used to inhibit intracellular triglyceride lipolysis through acting on metabolite-sensing Gi-protein-coupled GPR109A and subsequent PKA activation [36, 58, 61]. A study in rats suggested the brown adipocyte intracellular lipid lipolysis played a major role in thermogenesis in rodents [36]. It showed that the intracellular triglyceride lipolysis contributes to 84% of thermogenesis during an acute cold challenge (10°C, 2–6 hours) and 74% of thermogenesis during a chronic cold challenge (10°C, 21 days) [36]. The importance of the brown adipocyte intracellular triglyceridederived fatty acids was also reported in a human study [58]. In this study, administrating intracellular triglyceride lipolysis inhibitor NiAc significantly blunted BAT oxidative metabolism in cold-challenged young healthy humans (average 30 years

NiAc administration suppressed BAT intracellular triglyceride lipolysis by about 50% and BAT oxidative metabolism by 70%, despite of the increased blood flow in

One caveat from both the aforementioned human and rodent *in vivo* studies is that NiAc-mediated lipolysis inhibition took effect in both brown and white adipocytes in addition to other tissues, which makes it hard to delineate the contribution of lipolysis from each cell type. An *in vitro* study nicely confirmed the importance of intracellular lipolysis in brown adipocytes [2]. In cultured primary mouse brown adipocytes, the inhibition of both of the key lipolysis enzymes adipose triglyceride lipase (ATGL) and hormone-sensitive lipase (HSL) led to a 97% decrease in isopropanol-induced brown adipocyte respiration, indicating the imperative requirement of intracellular lipolysis in activated brown adipocytes [2]. These studies suggested that fatty acids liberated from the intracellular triglyceride storage serve as a critical fuel resource and contribute significantly to BAT thermogenesis in brown adipo-

Further studies in genetically manipulated mice showed similar results. The ATGL-knockout mouse presented defective triglyceride lipolysis function with increased BAT weight (8.5-fold), enlarged lipid droplet (20-fold), and decreased BAT explant lipid hydrolysis activity (−85%) [38]. The impaired triglyceride lipolysis activity in the *ATGL-knockout* mouse led to a defective thermogenic function. Upon a 5-hour 4°C challenge, mouse body temperature dropped to a critical low point at around 25°C [38]. Another study using mice with CGI-58 deficiency in both white and brown adipocytes also showed decreased BAT thermogenic function. The *adipose-CGI-58-knockout* mice were cold-sensitive, but only under fasted state [60]. The *HSL-null* mouse adipose tissue also presented defected triglyceride lipolysis function. Upon catecholamine-stimulation, *HSL-null* adipose tissue explants exhibited significantly reduced fatty acid and almost blunted glycerol release into the culture medium, parallel with diglycerides accumulation in both white and brown adipose tissue [56, 62]. However, it seems that HSL-mediated lipolysis function is not as critical as ATGL-mediated lipolysis function in brown adipocytes. An in vitro study showed that isopropanol-induced UCP1 activity was

). During a 3-hour cold challenge at 10°C,

#### *Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

*Cellular Metabolism and Related Disorders*

**2. Fatty acid metabolism in brown adipocytes**

acids serve as the activator for UCP1, which is a fatty acid/H+

thermogenic protein UCP1 in brown adipocytes [42, 43].

Thermogenic brown adipocyte possesses a high capacity for fatty acid β-oxidation that has been reported in both rodent and humans [4, 12, 14, 36]. Fatty

mediating proton flow and thermogenesis [37]. Fatty acids also serve as the main energy substrate mediating the uncoupling and thermogenic function in brown adipocytes [2, 36–41]. In addition, fatty acids can enhance brown adipocyte thermogenic capacity through the nuclear receptor peroxisome proliferator-activated receptors (PPARs), which are the master transcription regulators for the expression of genes involved in lipid metabolism, oxidative phosphorylation, and the key

Intracellular fatty acids are stored in the format of triglyceride in the heterogeneous multilocular lipid droplets in the brown adipocyte [4]. Upon the cold challenge, triglycerides stored in the brown adipocyte lipid droplet are lipolysed. Triglyceride lipolysis is a sequential process that involves different enzymes, resulting in the liberation of glycerol and fatty acids for heat production [44]. The lipid droplet is composed of triglycerides and cholesterol esters, which are surrounded by a monolayer of phospholipids [44]. Important proteins with regulatory and enzymatic functions, including perilipin and CGI58, coexist on the phospholipid monolayer to regulate the lipid trafficking and other functions of the lipid droplet [44–47]. Perilipin stabilizes the lipid droplet and prevents it from lipolysis under basal condition. Upon cold challenge, β-adrenergic stimulation leads to the activation of G-protein-coupled receptor (GPCR) and adenylate cyclase, which subsequently increases the cAMP level in brown adipocyte [48]. cAMP then activates protein kinase A (PKA), which phosphorylates perilipin at its serine residues [49–52]. The phosphorylated perilipin releases CGI-58, an adipose triglyceride lipase (ATGL)-activating protein. CGI-58, subsequently, binds and actives ATGL. Activated ATGL hydrolyzes triglycerides and generates free fatty acids and diglycerides [50, 53–55]. Upon the cold challenge, PKA also phosphorylates serine residues on another key lipolysis enzyme hormone sensitive lipase (HSL) [56]. Although HSL is capable of hydrolyzing triglycerides, diglycerides, monoglycerides,

symporter directly

adipocyte, energy generated from nutrients is initially stored as proton gradient membrane potential across the mitochondrial inner membrane, and then converted directly to thermo energy by the uncoupling protein 1 (UCP1)-mediated proton flow [4, 29]. In BAT, brown adipocytes are surrounded by abundant capillaries. The heat generated in the brown adipocyte mitochondria can be quickly distributed by the blood flow to maintain the steady core body temperature in mammals [4]. In addition to classical brown adipocytes, beige/brite adipocytes can be induced from the white adipocytes to conduct thermogenesis upon the cold challenge or catecholamine stimulation [26, 30, 31]. This process is termed as "browning" [26, 30, 31]. Thermogenesis in beige/brite adipocytes can also contribute to mammals' body temperature maintenance and whole body metabolism [26, 30, 31]. Beige/brite adipose tissue generates heat through both UCP1-independent thermogenesis, including calcium cycling in and out of endoplasmic reticulum, futile cycle between creatine and phosphocreatine, and UCP1-dependent thermogenesis in mitochondria [26, 30–33]. The energy metabolism is essential for the optimal UCP1-mediated mitochondrial thermogenic function of brown and beige/ brite adipocytes [4, 34, 35]. In this chapter, we discuss the importance of energy metabolism in maintaining brown adipocyte thermogenic function and the proceeding of targeting metabolic disorder through BAT activation in human studies.

**10**

and a broad array of other lipid substrate, it is the rate-limiting enzyme for hydrolyzing the diglycerides *in vivo* [56]. The phosphorylated HSL catalyzes the diglycerides and generates free fatty acids and monoglycerides [56]. As the last step of lipolysis, monoglycerides is hydrolyzed by monoacylglycerol lipase (MGL) to form free fatty acids and glycerol [57].

Given the importance of fatty acid in brown adipocyte thermogenic function, it is reasonable to predict that the deficiency of the key lipolysis enzymes and fatty acid transportation proteins in brown adipocytes can lead to a defected BAT thermogenic function. Human and rodent *in vivo* studies using both pharmacological and genetic approaches to manipulate triglyceride lipolysis processes were reported [36, 39, 58–60]. In the initial studies, nicotinic acid (NiAc) was used to inhibit intracellular triglyceride lipolysis through acting on metabolite-sensing Gi-protein-coupled GPR109A and subsequent PKA activation [36, 58, 61]. A study in rats suggested the brown adipocyte intracellular lipid lipolysis played a major role in thermogenesis in rodents [36]. It showed that the intracellular triglyceride lipolysis contributes to 84% of thermogenesis during an acute cold challenge (10°C, 2–6 hours) and 74% of thermogenesis during a chronic cold challenge (10°C, 21 days) [36]. The importance of the brown adipocyte intracellular triglyceridederived fatty acids was also reported in a human study [58]. In this study, administrating intracellular triglyceride lipolysis inhibitor NiAc significantly blunted BAT oxidative metabolism in cold-challenged young healthy humans (average 30 years of age with an average BMI of 24.5 kg/m2 ). During a 3-hour cold challenge at 10°C, NiAc administration suppressed BAT intracellular triglyceride lipolysis by about 50% and BAT oxidative metabolism by 70%, despite of the increased blood flow in the BAT [58].

One caveat from both the aforementioned human and rodent *in vivo* studies is that NiAc-mediated lipolysis inhibition took effect in both brown and white adipocytes in addition to other tissues, which makes it hard to delineate the contribution of lipolysis from each cell type. An *in vitro* study nicely confirmed the importance of intracellular lipolysis in brown adipocytes [2]. In cultured primary mouse brown adipocytes, the inhibition of both of the key lipolysis enzymes adipose triglyceride lipase (ATGL) and hormone-sensitive lipase (HSL) led to a 97% decrease in isopropanol-induced brown adipocyte respiration, indicating the imperative requirement of intracellular lipolysis in activated brown adipocytes [2]. These studies suggested that fatty acids liberated from the intracellular triglyceride storage serve as a critical fuel resource and contribute significantly to BAT thermogenesis in brown adipocytes upon activation.

Further studies in genetically manipulated mice showed similar results. The ATGL-knockout mouse presented defective triglyceride lipolysis function with increased BAT weight (8.5-fold), enlarged lipid droplet (20-fold), and decreased BAT explant lipid hydrolysis activity (−85%) [38]. The impaired triglyceride lipolysis activity in the *ATGL-knockout* mouse led to a defective thermogenic function. Upon a 5-hour 4°C challenge, mouse body temperature dropped to a critical low point at around 25°C [38]. Another study using mice with CGI-58 deficiency in both white and brown adipocytes also showed decreased BAT thermogenic function. The *adipose-CGI-58-knockout* mice were cold-sensitive, but only under fasted state [60]. The *HSL-null* mouse adipose tissue also presented defected triglyceride lipolysis function. Upon catecholamine-stimulation, *HSL-null* adipose tissue explants exhibited significantly reduced fatty acid and almost blunted glycerol release into the culture medium, parallel with diglycerides accumulation in both white and brown adipose tissue [56, 62]. However, it seems that HSL-mediated lipolysis function is not as critical as ATGL-mediated lipolysis function in brown adipocytes. An in vitro study showed that isopropanol-induced UCP1 activity was

largely dependent on ATGL function (80%) compared to HSL function (35%) in cultured primary brown adipocytes, and the combined inhibition of both ATGL and HSL functions led to an almost complete block of UCP1-mediated thermogenic function (97%) [2].

These studies highlight the cardinal role of intracellular triglyceride liberation in the brown adipocyte thermogenic function, but raise the question if the brown adipocyte intracellular lipolysis is essential for BAT to maintain the adaptive thermogenesis *in vivo*. Interestingly, recent studies using *UCP1-Cre-*mediated knockout of either ATGL or CGI-58 gene in mice showed an unexpected insignificant impact of brown adipocyte intracellular lipolysis and suggested that the circulating fatty acids mobilized from peripheral tissues may play more important roles in BAT thermogenesis in mice [59, 60]. These studies demonstrated that the loss of either ATGL or CGI58-mediated triglycerides lipolysis in brown adipocytes did not compromise the cold challenge-induced thermogenic response in mice [59, 60]. The phenotypic discrepancy between the systemic and BAT-specific *ATGL* and *CGI-58* knockout mice brings very important insights to *in vivo* BAT fatty acid metabolism, and indicates that the brown adipocyte intracellular lipolysis is not the only energy resource for *in vivo* adaptive thermogenesis during cold challenge. Although it is not clear about the exact partition of intracellular and circulating fatty acids contribution to the BAT thermogenic function, it is reasonable to speculate that the core body temperature maintenance is vital for mammals to maintain their optimal function during the cold challenge, when the intracellular lipolysis function is impaired or insufficient, circulating energy substrates from other metabolic tissues, that is, white adipose tissue and liver, are mobilized to provide energy substrates for BAT-mediated adaptive thermogenesis to maintain the adequate core body temperature and ensure the optimal functionality of mammals.

Long-chain fatty acids (LCFAs) are the most abundant format of fatty acid energy substrate in mammals [63–66]. The liberated LCFAs from intracellular lipid droplet are facilitated and transported to mitochondrion and nucleus by fatty acid-binding proteins (FABPs) to conduct their functions [67]. Of the six FABP isoforms, FABP4 (also termed adipocyte p2) and FABP5 are the major FABP isoforms in brown adipocytes [68–74]. Mice mutated in both *FABP4* and *FABP5* gene developed severe hypothermia during fasting after an acute cold challenge (1–3 hours), indicating that fatty acids transportation plays an indispensable role in BAT thermogenic function [74].

The LCFAs-mediated mitochondrial oxidation and UCP-1 activation function require sequential carnitine acyltransferases in order to translocate the LCFA into mitochondrial matrix. Carnitine palmitoyltransferase 1 (CPT1), located on the mitochondrial outer membrane, is the rate-limiting enzyme that mediates LCFAs inward translocation into mitochondrial matrix [63–66]. CPT1 exists in tissuespecific isoforms, and CPT1b is the major isoform expressed in brown adipocytes [75–77]. Mouse embryos-carried homozygous knockout of *Cpt1b* gene were lethal before embryonic day 9.5–11.5 and a normal percentage of *CPT1b+/−* mice was born from the *CPT1b+/−* and wild type breeding pairs [78]. However, more than 50% of the *CPT1b+/−* pups were lost before weaning [78]. The detailed experiment showed that ∼7% *CPT1b+/−* mice developed fatal hypothermia following a 3-hour cold challenge (4°C) and ∼52% *CPT1b+/−* mice developed fatal hypothermia following a 6-hour cold challenge (4°C), indicating the critical contribution of CPT1b-mediated LCFAs transportation and thermogenesis during the cold challenge in infant/young mice [78]. Carnitine palmitoyltransferase 2 (CPT2) is located on mitochondrial inner membrane to further medicate LCFAs translocation into mitochondrial matrix. In line with the study using the *CPT1b-*deficient mice, an *adipose-specific CPT2 knockout* mice also presented hypothermic phenotype after 3 hours cold

**13**

an oral 3

*Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

tion during the cold challenge.

thermogenic function in mitochondria.

challenge (4°C) and decreased LCFAs oxidation in isolated MEF cells [39, 79]. These studies indicate that fatty acid transportation is critical for BAT thermogenic func-

The intramitochondrial LCFAs contribute to the thermogenesis through UCP-1 activation and β-oxidation. A detailed study confirmed that mice with impaired fatty acid β-oxidation are cold-intolerant [80]. The acyl-CoA dehydrogenases, which catalyze the initial steps of fatty acid β-oxidation, are composed of a group of enzymes, including very long-chain acyl CoA dehydrogenase (VLCAD), longchain acyl CoA dehydrogenase (LCAD), and short-chain acyl CoA dehydrogenase (SCAD) [81, 82]. A detailed experiment showed that fetal hypothermia was presented in all of the homozygous knockout mice (*VLCAD−/−, LCAD−/−*, and *SCAD−/−*) after 1-hour 4°C challenge [80]. Although the mice with single heterozygous of VLCAD, LCAD, and SCAD genes were cold tolerate; more than 30% of mice with double heterozygous *VLCAD+/−//LCAD+/− or LCAD+/−//SCAD+/−* and triple heterozygous *VLCAD+/−//LCAD+/−//SCAD+/−* combinations developed hypothermia upon the cold challenge [80], indicating the essential role of the LCFAs-mediated

In summary, these studies highlight the importance of the brown adipocyte lipolysis and the liberated intracellular fatty acid transportation and oxidation during thermogenesis. In addition, these studies indicate that brown adipocytes not only use the intracellular lipid storage, but also the circulating energy substrate to maintain the critical thermogenic function for the organismal survival and optimal function in mammals [2, 12, 38, 44, 56, 59, 60, 80]. In modern days, BAT's ability to metabolize fatty acids mobilized from other peripheral storages, including white adipose tissue and liver, makes it a good potential therapeutic target in humans. Studies have shown that BAT mediates significant plasma lipid clearance during the cold challenge in rodent and humans under both physiological and pathological conditions [83–85]. One study showed that the activated BAT is involved in the basal and postprandial triglyceride metabolism in rodents [83]. In this study, compared to mice kept at 22°C, mice kept at 4°C had significantly lower triglyceride-rich lipoproteins (TRLs)-triglyceride concentration [83]. The study also showed that the activated BAT was involved in the post-prandial triglyceride metabolic process, evidenced by

H-triolein tolerance test. Under cold challenge condition, the BAT 3

uptake was significantly higher than that of either liver or skeletal muscle, suggesting the significant triglyceride/triglyceride-rich lipoprotein metabolism in activated BAT. For the triglyceride clearance, circulating triglycerides rose to a peak at 2 hours postprandial and declined subsequently in mice kept at 22°C. In contrast, the triglyceride level reminded persistently low in mice kept at 4°C, suggesting that the BAT possesses a high postprandial triglyceride clearance function [83]. Most interestingly, the cold challenge increased the uptake of radio-labeled lipoprotein in BAT and reduced the uptake in liver [83]. The cold challenge-induced lipid clearance shift suggests that BAT can be targeted for lipid metabolism *in vivo*. In a pathological setting, the genetically manipulated *Apoa5−/−* mice with severe hyperlipidemia were studied. A 4–24 hours 4°C challenge corrected *Apoa5−/−* mouse plasma lipid concentration to the values comparable to wild-type mice, indicating the significant impact of BAT on

whole body lipid metabolism in the rodent under the pathological condition [83, 86, 87]. Other studies also showed that the BAT preferentially uptook plasma triglycerides through peripheral tissue lipolysis, and the selective fatty acids uptake from triglyceride-rich lipoprotein ameliorated hyperlipidemia in rodents [84, 85]. Although the studies in rodent strongly support the importance of BAT in lipid clearance, the significance of BAT in human lipid metabolism is still unclear. The contribution of activated BAT in human body was studied using a dietary radiolabeled LCFAs tracer 14(R,S)-[18F]-fluoro-6-thia-heptadecanoic acid (18FTHA) [88].

H-triolein

#### *Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

*Cellular Metabolism and Related Disorders*

function (97%) [2].

largely dependent on ATGL function (80%) compared to HSL function (35%) in cultured primary brown adipocytes, and the combined inhibition of both ATGL and HSL functions led to an almost complete block of UCP1-mediated thermogenic

These studies highlight the cardinal role of intracellular triglyceride liberation in the brown adipocyte thermogenic function, but raise the question if the brown adipocyte intracellular lipolysis is essential for BAT to maintain the adaptive thermogenesis *in vivo*. Interestingly, recent studies using *UCP1-Cre-*mediated knockout of either ATGL or CGI-58 gene in mice showed an unexpected insignificant impact of brown adipocyte intracellular lipolysis and suggested that the circulating fatty acids mobilized from peripheral tissues may play more important roles in BAT thermogenesis in mice [59, 60]. These studies demonstrated that the loss of either ATGL or CGI58-mediated triglycerides lipolysis in brown adipocytes did not

compromise the cold challenge-induced thermogenic response in mice [59, 60]. The phenotypic discrepancy between the systemic and BAT-specific *ATGL* and *CGI-58* knockout mice brings very important insights to *in vivo* BAT fatty acid metabolism, and indicates that the brown adipocyte intracellular lipolysis is not the only energy resource for *in vivo* adaptive thermogenesis during cold challenge. Although it is not clear about the exact partition of intracellular and circulating fatty acids contribution to the BAT thermogenic function, it is reasonable to speculate that the core body temperature maintenance is vital for mammals to maintain their optimal function during the cold challenge, when the intracellular lipolysis function is impaired or insufficient, circulating energy substrates from other metabolic tissues, that is, white adipose tissue and liver, are mobilized to provide energy substrates for BAT-mediated adaptive thermogenesis to maintain the adequate core body tempera-

Long-chain fatty acids (LCFAs) are the most abundant format of fatty acid energy substrate in mammals [63–66]. The liberated LCFAs from intracellular lipid droplet are facilitated and transported to mitochondrion and nucleus by fatty acid-binding proteins (FABPs) to conduct their functions [67]. Of the six FABP isoforms, FABP4 (also termed adipocyte p2) and FABP5 are the major FABP isoforms in brown adipocytes [68–74]. Mice mutated in both *FABP4* and *FABP5* gene developed severe hypothermia during fasting after an acute cold challenge (1–3 hours), indicating that fatty acids transportation plays an indispensable role in

The LCFAs-mediated mitochondrial oxidation and UCP-1 activation function require sequential carnitine acyltransferases in order to translocate the LCFA into mitochondrial matrix. Carnitine palmitoyltransferase 1 (CPT1), located on the mitochondrial outer membrane, is the rate-limiting enzyme that mediates LCFAs inward translocation into mitochondrial matrix [63–66]. CPT1 exists in tissuespecific isoforms, and CPT1b is the major isoform expressed in brown adipocytes [75–77]. Mouse embryos-carried homozygous knockout of *Cpt1b* gene were lethal before embryonic day 9.5–11.5 and a normal percentage of *CPT1b+/−* mice was born from the *CPT1b+/−* and wild type breeding pairs [78]. However, more than 50% of the *CPT1b+/−* pups were lost before weaning [78]. The detailed experiment showed that ∼7% *CPT1b+/−* mice developed fatal hypothermia following a 3-hour cold challenge (4°C) and ∼52% *CPT1b+/−* mice developed fatal hypothermia following a 6-hour cold challenge (4°C), indicating the critical contribution of CPT1b-mediated LCFAs transportation and thermogenesis during the cold challenge in infant/young mice [78]. Carnitine palmitoyltransferase 2 (CPT2) is located on mitochondrial inner membrane to further medicate LCFAs translocation into mitochondrial matrix. In line with the study using the *CPT1b-*deficient mice, an *adipose-specific CPT2 knockout* mice also presented hypothermic phenotype after 3 hours cold

ture and ensure the optimal functionality of mammals.

BAT thermogenic function [74].

**12**

challenge (4°C) and decreased LCFAs oxidation in isolated MEF cells [39, 79]. These studies indicate that fatty acid transportation is critical for BAT thermogenic function during the cold challenge.

The intramitochondrial LCFAs contribute to the thermogenesis through UCP-1 activation and β-oxidation. A detailed study confirmed that mice with impaired fatty acid β-oxidation are cold-intolerant [80]. The acyl-CoA dehydrogenases, which catalyze the initial steps of fatty acid β-oxidation, are composed of a group of enzymes, including very long-chain acyl CoA dehydrogenase (VLCAD), longchain acyl CoA dehydrogenase (LCAD), and short-chain acyl CoA dehydrogenase (SCAD) [81, 82]. A detailed experiment showed that fetal hypothermia was presented in all of the homozygous knockout mice (*VLCAD−/−, LCAD−/−*, and *SCAD−/−*) after 1-hour 4°C challenge [80]. Although the mice with single heterozygous of VLCAD, LCAD, and SCAD genes were cold tolerate; more than 30% of mice with double heterozygous *VLCAD+/−//LCAD+/− or LCAD+/−//SCAD+/−* and triple heterozygous *VLCAD+/−//LCAD+/−//SCAD+/−* combinations developed hypothermia upon the cold challenge [80], indicating the essential role of the LCFAs-mediated thermogenic function in mitochondria.

In summary, these studies highlight the importance of the brown adipocyte lipolysis and the liberated intracellular fatty acid transportation and oxidation during thermogenesis. In addition, these studies indicate that brown adipocytes not only use the intracellular lipid storage, but also the circulating energy substrate to maintain the critical thermogenic function for the organismal survival and optimal function in mammals [2, 12, 38, 44, 56, 59, 60, 80]. In modern days, BAT's ability to metabolize fatty acids mobilized from other peripheral storages, including white adipose tissue and liver, makes it a good potential therapeutic target in humans. Studies have shown that BAT mediates significant plasma lipid clearance during the cold challenge in rodent and humans under both physiological and pathological conditions [83–85].

One study showed that the activated BAT is involved in the basal and postprandial triglyceride metabolism in rodents [83]. In this study, compared to mice kept at 22°C, mice kept at 4°C had significantly lower triglyceride-rich lipoproteins (TRLs)-triglyceride concentration [83]. The study also showed that the activated BAT was involved in the post-prandial triglyceride metabolic process, evidenced by an oral 3 H-triolein tolerance test. Under cold challenge condition, the BAT 3 H-triolein uptake was significantly higher than that of either liver or skeletal muscle, suggesting the significant triglyceride/triglyceride-rich lipoprotein metabolism in activated BAT. For the triglyceride clearance, circulating triglycerides rose to a peak at 2 hours postprandial and declined subsequently in mice kept at 22°C. In contrast, the triglyceride level reminded persistently low in mice kept at 4°C, suggesting that the BAT possesses a high postprandial triglyceride clearance function [83]. Most interestingly, the cold challenge increased the uptake of radio-labeled lipoprotein in BAT and reduced the uptake in liver [83]. The cold challenge-induced lipid clearance shift suggests that BAT can be targeted for lipid metabolism *in vivo*. In a pathological setting, the genetically manipulated *Apoa5−/−* mice with severe hyperlipidemia were studied. A 4–24 hours 4°C challenge corrected *Apoa5−/−* mouse plasma lipid concentration to the values comparable to wild-type mice, indicating the significant impact of BAT on whole body lipid metabolism in the rodent under the pathological condition [83, 86, 87]. Other studies also showed that the BAT preferentially uptook plasma triglycerides through peripheral tissue lipolysis, and the selective fatty acids uptake from triglyceride-rich lipoprotein ameliorated hyperlipidemia in rodents [84, 85].

Although the studies in rodent strongly support the importance of BAT in lipid clearance, the significance of BAT in human lipid metabolism is still unclear. The contribution of activated BAT in human body was studied using a dietary radiolabeled LCFAs tracer 14(R,S)-[18F]-fluoro-6-thia-heptadecanoic acid (18FTHA) [88]. This study showed that a 4-hour mild cold-challenge at 18°C significantly increased dietary fatty acids distribution in BAT in humans [88]. However, given the relative small volume of BAT tissue, the dietary fatty acids clearance contribution from the BAT is less significant compared to other organs including heart, liver, white adipose tissue, or skeletal muscles, and only contributed to ~0.3% of total body dietary fatty acids clearance upon the mild cold challenge [88]. Similarly, another study using fatty acid tracer 18F-fluoro-thiaheptadecanoic acid (18FTHA) showed that a 3-hour cold challenge at 18°C led to four times higher radio-labeled tracer uptake and ~80% metabolic rate increase in the BAT, contributing <1% of total fatty acids clearance rate in human subjects [12]. Nevertheless, these experiments suggest that BAT not only exists, but also is functionally active and can contribute to systemic metabolism in humans. One possible reason for the relatively low BAT contribution in fatty acids metabolism in humans could be due to the short cold challenge time and mild cold challenge conditions. It is possible that during the acute cold challenge, intracellular lipid lipolysis serves as the main energy resource so that it ameliorates the clearance of circulating TRLs or fatty acids.

The circulating TRLs or fatty acids are transported into brown adipocyte by a group of proteins, including lipoprotein lipase (LPL) and fatty acid transport proteins [83, 89–93]. LPL is a multifunctional protein produced by many tissues, including the adipose tissue [94]. LPL serves as a rate-limiting enzyme mediating extracellular lipolysis [94]. It hydrolyzes triglycerides into lipid-rich proteins into fatty acids and monoacylglycerol. It also mediates the cellular uptake of triglyceride and other lipids [94]. Studies have shown that cold challenge or catecholaminestimulation induce the expression and activity of LPL in brown adipocytes through a cAMP-mediated mechanism [89–91]. After activation, LPL is released from the brown adipocyte, transferred to the capillary endothelium lumen, and serves as the anchor between the endothelium cell surface and the TRLs [95–97]. Next, the LCFAs liberated from LPL channel into brown adipocyte for thermogenic function [98, 99]. A study indicated that the local LPL activity is required for TRLs uptake into the BAT [83]. In this study, it is shown that either LPL-specific inhibitor tetrahydrolipstatin pretreatment or releasing LPL from endothelium by heparin significantly blocked the uptake of TRL and fatty acids in BAT [83].

The liberated LCFAs in circulation can be transported into cells and activated by both transmembrane fatty acid transporter proteins (FATPs) and scavenger receptor CD36 [92, 93]. FATPs are composed by a family of six proteins mediating circulating LCFAs uptake and distribution in cells [83, 92, 93, 100]. Among the six isoforms, FATP1 (SLC27A1) is the major isoform in brown and white adipose tissue [83, 92, 93, 100]. Studies showed that postprandial lipid uptake is highly dependent on the adipose tissue FATP1 [101]. The FATP1-knockout mice have decreased lipid uptake in adipose tissue and a compensatory lipid redistribution in liver and heart, where FATP1-mediated lipid uptake function is not required under normal conditions [101]. The cold challenge can induce FATP1 expression in BAT [100]. In line with this, the isolated FATP1-null brown adipocytes showed significantly less fatty acids uptake upon catecholamine stimulation [100]. *In vivo* studies showed that in cold challenged FATP1-knockout mice, BAT triglyceride storage was decreased and circulating serum free fatty acids was increased, indicating the importance of FATP1-mediated fatty acid uptake in BAT [100]. The LCFAs uptake can also be mediated by the transmembrane class B scavenger receptor CD36 [83, 102, 103]. Upon the cold challenge, CD36 is significantly upregulated in adipocytes [83, 102]. Other studies using *CD36−/−* mice clearly indicated the importance of CD36-mediated LCFAs transportation during thermogenesis. Around 60% of *Cd36−/−* mice died during a 24-hour cold challenge, which is paralleled with drastically increased plasma free fatty acids concentration [83, 102]. A recent

**15**

*Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

**3. Glucose metabolism in BAT**

challenge [4, 9–12, 25, 26, 107–112].

disorders.

the BAT mitochondrial thermogenic function [104].

study indicated that CD36-medicated coenzyme Q (CoQ ) uptake is required for

In summary, fatty acid is indispensable for the brown adipocyte thermogenic function. Fatty acid mediates brown adipocyte thermogenesis through mitochondrial β-oxidation, UCP1-activation, and fatty acid-mediated thermogenic gene regulation. Both the intracellular fatty acids from brown adipocyte lipolysis and the liberated fatty acids from peripheral tissues play essential roles mediating the critical BAT adaptive thermogenic function to main the adequate core body temperature in mammals. In modern days, activating BAT thermogenic function to increase fatty acids uptake and utilization may offer new therapeutics to treat human metabolic

The radio-labeled glucose analog 18F-fluorodeoxyglucose (18F-FDG), in combination with positron emission tomography (PET) and computed tomography (CT), provides a reliable method for the *in vivo* tissue glucose uptake study [105, 106]. Based on studies using this method, it is now well-recognized that BAT in both rodents and humans possesses a significant glucose uptake capacity upon the cold

Glucose uptake is regulated in brown adipocytes. *In vivo* studies showed that cold challenge significantly enhanced insulin sensitivity and subsequent glucose uptake in the BAT [9, 10, 25, 26, 107, 112, 113]. Interestingly, other studies showed that starved rats with low insulin level also had increased BAT glucose uptake during the cold challenge, indicating that the enhanced glucose uptake is not completely insulin-dependent [114]. Additional studies also indicated that the enhanced glucose uptake in brown adipocytes can be mediated by different pathways in addition to insulin stimulation, including β-adrenergic receptor activation, AMPactivated protein kinase (AMPK) activation, and mTOR activation [115–117]. The importance of glucose metabolism in BAT is a long-standing question. Glucose transporters, which facilitate glucose across the cell plasma membrane is the first rate-limiting step of the glucose metabolism [34, 118, 119]. Intracellular glucose is subsequently phosphorylated to glucose-6-phosphate (G6P) by the enzyme hexokinase (HK). Glucose-6-phosphate serves as the substrate into different pathways, including glycolysis, glycogen synthesis, and the pentose phosphate pathway (PPP) [34, 118, 119]. Glycolysis breaks down glucose to pyruvate to generate small amount of ATP and NADH [34, 118, 119]. The pyruvate is then transported into mitochondria for oxidation and energy production. Under hypoxia

condition, pyruvate is disposed in the form of lactate [34, 118, 119].

Early study indicates that glucose and its metabolites contribute to the brown adipocyte thermogenesis by showing that catecholamine-induced glucose uptake was decreased when mitochondrial β-oxidation was inhibited in brown adipocytes [120]. Other studies in brown adipocyte glucose transportation also indicate the importance of the glucose in brown adipocyte metabolism [6, 112, 121, 122]. Both glucose transporter 1 (Glut1) and glucose transporter 4 (Glut4) are abundantly expressed in brown adipocytes and the insulin sensitive Glut4 is the major isoform [112, 122]. The *in vitro* study showed that knock-down of Glut1 and/or Glut4 gene in cultured brown adipocytes impaired the catecholamine-induced cell oxygen consumption by 30–50% [6, 121]. Other studies indicate the importance of glycolysis in brown adipocyte thermogenesis [115, 123, 124]. An *in vitro* study showed that the knockdown of two glycolysis enzymes, HK2 or pyruvate kinase M (PKM), decreased glucose uptake and catecholamine-induced cell oxygen consumption by 67% or

#### *Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

*Cellular Metabolism and Related Disorders*

of circulating TRLs or fatty acids.

This study showed that a 4-hour mild cold-challenge at 18°C significantly increased dietary fatty acids distribution in BAT in humans [88]. However, given the relative small volume of BAT tissue, the dietary fatty acids clearance contribution from the BAT is less significant compared to other organs including heart, liver, white adipose tissue, or skeletal muscles, and only contributed to ~0.3% of total body dietary fatty acids clearance upon the mild cold challenge [88]. Similarly, another study using fatty acid tracer 18F-fluoro-thiaheptadecanoic acid (18FTHA) showed that a 3-hour cold challenge at 18°C led to four times higher radio-labeled tracer uptake and ~80% metabolic rate increase in the BAT, contributing <1% of total fatty acids clearance rate in human subjects [12]. Nevertheless, these experiments suggest that BAT not only exists, but also is functionally active and can contribute to systemic metabolism in humans. One possible reason for the relatively low BAT contribution in fatty acids metabolism in humans could be due to the short cold challenge time and mild cold challenge conditions. It is possible that during the acute cold challenge, intracellular lipid lipolysis serves as the main energy resource so that it ameliorates the clearance

The circulating TRLs or fatty acids are transported into brown adipocyte by a group of proteins, including lipoprotein lipase (LPL) and fatty acid transport proteins [83, 89–93]. LPL is a multifunctional protein produced by many tissues, including the adipose tissue [94]. LPL serves as a rate-limiting enzyme mediating extracellular lipolysis [94]. It hydrolyzes triglycerides into lipid-rich proteins into fatty acids and monoacylglycerol. It also mediates the cellular uptake of triglyceride and other lipids [94]. Studies have shown that cold challenge or catecholaminestimulation induce the expression and activity of LPL in brown adipocytes through a cAMP-mediated mechanism [89–91]. After activation, LPL is released from the brown adipocyte, transferred to the capillary endothelium lumen, and serves as the anchor between the endothelium cell surface and the TRLs [95–97]. Next, the LCFAs liberated from LPL channel into brown adipocyte for thermogenic function [98, 99]. A study indicated that the local LPL activity is required for TRLs uptake into the BAT [83]. In this study, it is shown that either LPL-specific inhibitor tetrahydrolipstatin pretreatment or releasing LPL from endothelium by heparin

significantly blocked the uptake of TRL and fatty acids in BAT [83].

The liberated LCFAs in circulation can be transported into cells and activated by both transmembrane fatty acid transporter proteins (FATPs) and scavenger receptor CD36 [92, 93]. FATPs are composed by a family of six proteins mediating circulating LCFAs uptake and distribution in cells [83, 92, 93, 100]. Among the six isoforms, FATP1 (SLC27A1) is the major isoform in brown and white adipose tissue [83, 92, 93, 100]. Studies showed that postprandial lipid uptake is highly dependent on the adipose tissue FATP1 [101]. The FATP1-knockout mice have decreased lipid uptake in adipose tissue and a compensatory lipid redistribution in liver and heart, where FATP1-mediated lipid uptake function is not required under normal conditions [101]. The cold challenge can induce FATP1 expression in BAT [100]. In line with this, the isolated FATP1-null brown adipocytes showed significantly less fatty acids uptake upon catecholamine stimulation [100]. *In vivo* studies showed that in cold challenged FATP1-knockout mice, BAT triglyceride storage was decreased and circulating serum free fatty acids was increased, indicating the importance of FATP1-mediated fatty acid uptake in BAT [100]. The LCFAs uptake can also be mediated by the transmembrane class B scavenger receptor CD36 [83, 102, 103]. Upon the cold challenge, CD36 is significantly upregulated in adipocytes [83, 102]. Other studies using *CD36−/−* mice clearly indicated the importance of CD36-mediated LCFAs transportation during thermogenesis. Around 60% of *Cd36−/−* mice died during a 24-hour cold challenge, which is paralleled with drastically increased plasma free fatty acids concentration [83, 102]. A recent

**14**

study indicated that CD36-medicated coenzyme Q (CoQ ) uptake is required for the BAT mitochondrial thermogenic function [104].

In summary, fatty acid is indispensable for the brown adipocyte thermogenic function. Fatty acid mediates brown adipocyte thermogenesis through mitochondrial β-oxidation, UCP1-activation, and fatty acid-mediated thermogenic gene regulation. Both the intracellular fatty acids from brown adipocyte lipolysis and the liberated fatty acids from peripheral tissues play essential roles mediating the critical BAT adaptive thermogenic function to main the adequate core body temperature in mammals. In modern days, activating BAT thermogenic function to increase fatty acids uptake and utilization may offer new therapeutics to treat human metabolic disorders.

#### **3. Glucose metabolism in BAT**

The radio-labeled glucose analog 18F-fluorodeoxyglucose (18F-FDG), in combination with positron emission tomography (PET) and computed tomography (CT), provides a reliable method for the *in vivo* tissue glucose uptake study [105, 106]. Based on studies using this method, it is now well-recognized that BAT in both rodents and humans possesses a significant glucose uptake capacity upon the cold challenge [4, 9–12, 25, 26, 107–112].

Glucose uptake is regulated in brown adipocytes. *In vivo* studies showed that cold challenge significantly enhanced insulin sensitivity and subsequent glucose uptake in the BAT [9, 10, 25, 26, 107, 112, 113]. Interestingly, other studies showed that starved rats with low insulin level also had increased BAT glucose uptake during the cold challenge, indicating that the enhanced glucose uptake is not completely insulin-dependent [114]. Additional studies also indicated that the enhanced glucose uptake in brown adipocytes can be mediated by different pathways in addition to insulin stimulation, including β-adrenergic receptor activation, AMPactivated protein kinase (AMPK) activation, and mTOR activation [115–117].

The importance of glucose metabolism in BAT is a long-standing question. Glucose transporters, which facilitate glucose across the cell plasma membrane is the first rate-limiting step of the glucose metabolism [34, 118, 119]. Intracellular glucose is subsequently phosphorylated to glucose-6-phosphate (G6P) by the enzyme hexokinase (HK). Glucose-6-phosphate serves as the substrate into different pathways, including glycolysis, glycogen synthesis, and the pentose phosphate pathway (PPP) [34, 118, 119]. Glycolysis breaks down glucose to pyruvate to generate small amount of ATP and NADH [34, 118, 119]. The pyruvate is then transported into mitochondria for oxidation and energy production. Under hypoxia condition, pyruvate is disposed in the form of lactate [34, 118, 119].

Early study indicates that glucose and its metabolites contribute to the brown adipocyte thermogenesis by showing that catecholamine-induced glucose uptake was decreased when mitochondrial β-oxidation was inhibited in brown adipocytes [120]. Other studies in brown adipocyte glucose transportation also indicate the importance of the glucose in brown adipocyte metabolism [6, 112, 121, 122]. Both glucose transporter 1 (Glut1) and glucose transporter 4 (Glut4) are abundantly expressed in brown adipocytes and the insulin sensitive Glut4 is the major isoform [112, 122]. The *in vitro* study showed that knock-down of Glut1 and/or Glut4 gene in cultured brown adipocytes impaired the catecholamine-induced cell oxygen consumption by 30–50% [6, 121]. Other studies indicate the importance of glycolysis in brown adipocyte thermogenesis [115, 123, 124]. An *in vitro* study showed that the knockdown of two glycolysis enzymes, HK2 or pyruvate kinase M (PKM), decreased glucose uptake and catecholamine-induced cell oxygen consumption by 67% or

34% respectively, in brown adipocytes [6]. These studies suggested the importance of glucose metabolism in brown adipocyte function. However, another detailed study indicated that glucose oxidation does not play a major role in brown adipocyte metabolism and thermogenesis, by showing that the rate of 14CO2 formation from the 14C glucose was relatively small compared with the maximum rate of oxygen consumption in activated brown adipocytes [125]. Studies using radio-labeled glucose also suggested that glucose uptake was only sufficient to fuel maximally ~15% of the thermogenic capacity in activated rodent brown adipocytes, suggesting that the significantly upregulated glucose uptake in activated brown adipocytes is disassociated with the relative low glucose-mediated thermogenic capacity [123, 125, 126]. A more recent study indicates that the brown adipocyte energy production from glucose depends on the state of the cell: glucose and fatty acid contribute equally to brown adipocyte oxygen consumption under the basal condition; upon catecholamineactivation, oxygen consumption is mainly fueled by fatty acids [6].

The dissociation between high glucose uptake and low glucose-mediated thermogenesis in activated BAT raises an important question: what is the function of the intracellular glucose in brown adipocytes? The importance of glucose in the *de novo* lipogenesis in brown adipocytes has been reported [127–129]. Studies indicate that glucose uptake is an independent process of thermogenesis in both cold-challenged and catecholamine-activated BAT, which further supports that glucose uptake might play other role as energy substrate in activated brown adipocytes [116, 117, 130–132]. One detailed *in vitro* study using rat brown adipocytes showed that norepinephrine significantly enhanced glucose oxidation by sevenfold, while it also inhibited lipogenesis at the same time. On the other hand, insulin stimulation increased the lipogenesis by sevenfold in brown adipocytes whereas glucose oxidation remained very low. Most interestingly, the addition of insulin to the norepinephrine only potentiated the enhanced glucose oxidation, but do not enhance the lipogenesis [115]. On the other hand, another study showed that brown adipocytes converted a greater proportion of metabolized glucose into lactate and pyruvate, but only a smaller proportion into fatty acids through insulin-mediated pathway [125]. These studies suggest that glucose metabolism is involved in two different states in brown adipocytes, the thermogenic state and nonthermogenic state. It is plausible that during the thermogenic state, glucose contributes to both the energy production and lipogenesis; and during the nonthermogenic state, glucose contributes mainly to the lipogenesis and energy storage process in brown adipocytes, which explains the disassociation of glucose uptake and glucose metabolism in brown adipocytes. In addition to glucose, other energy substrates, including glycerol and amino acid (glutamate), might also contribute to BAT metabolism and thermogenesis in human and rodent brown adipocytes. However, the relative contribution and partition of these energy substrates are unclear [132–135].

#### **4. Energy storage in BAT**

The cold challenge not only enhances catabolic processes mediating energy substrate metabolism and heat generation, but also induces anabolic processes mediating fatty acid synthesis and lipogenesis, as well as glycogenesis [36, 111, 136–139].

Glucose can be stored as glycogen in brown adipocytes [36, 139]. Studies showed that glycogen synthases (GStot) and uridine diphosphate glucose pyrophosphorylase (Udgp), which mediates glycogenesis, were upregulated upon the cold challenge in BAT [36, 139]. Interestingly, glycogen hydrolysis enzyme, glycogen phosphorylase (Pygl), was also upregulated after cold challenge [36]. Although cold-challenge upregulates both glycogen synthesis and degradation, it is reported

**17**

energy homeostasis.

*Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

resource in brown adipocytes.

that the stored glycogen is used up shortly after the cold challenge [139]. Indicating that glycogen is not an efficient format for energy storage and a sustainable energy

The glucose uptaken by brown adipocyte can also be converted to fatty acid through the *de novo* lipogenesis. Carbohydrate response element-binding protein (ChREBP) is the major transcription factor mediating fatty acid synthesis in adipocytes. There are two isoforms of ChREBP, ChREBPα and ChREBPβ. ChREBPα is abundantly expressed in BAT from rodents and humans [137, 140, 141]. *De novo* lipogenesis involves a series of enzymes mediating the sequential reactions converting glucose-derived citrate into fatty acids, including ATP-citrate lyase (ACLY), acetyl-CoA carboxylases 1 (ACC1), fatty acid synthase (FASN), and stearoyl-CoA desaturase-1 (SCD1). ChREBPα, coordinates with another transcription factor Max-like protein (MLX), directly binds to the carbohydrate response elements (ChoRE) and upregulates these *de novo* lipogenic genes expressions. ChREBPα can also increase the expression of ChREBPβ to further activate the *de novo* lipogenesis enzymes [142, 143]. It has been reported that lipogenic genes and AKT2-ChREBP pathways are upregulated to optimize fuel storage and thermogenesis upon cold stimulation in BAT [136]. In accordance with these studies, *ChREBP−/−* mice presented less BAT weight [142], and adipose-specific *ChREBP-knockout* mice had decreased carbohydrate-induced lipogenesis in BAT [144]. These studies indicate that ChREBP plays important roles in brown adipocyte's *de novo* lipogenesis and energy storage.

Sterol regulatory element-binding protein-1 (SREBP-1) is another transcription factor mediating the *de novo* lipogenesis [145]. Of the three different SREBP isoforms, SREBP1c is more abundant and SREBP1a is less abundant in the adipose tissue [137, 145, 146]. *In vitro* study showed that SREBP1c is sufficient to regulate lipogenic enzymes in cultured adipocytes [147]. In the adipose-specific *ap2- SREBP1c* transgenic mice, lipogenic enzyme Acc1, FASN and Scd1 expression as well as fatty acids synthesis rate were significantly upregulated in brown adipocytes [148]. In line with this study, *ap2-SREBP1a* transgenic mice also developed adipose tissue hypertrophy in accordance with an increased lipogenic enzyme profile and enhance *de novo* lipogenesis in the BAT [148]. However, some *in vivo* studies indicated that SREBP1c's role in the *de novo* lipogenesis in adipocytes is dispensable, as evidenced by *SREBP1-knockout* mice have normal lipogenic enzymes gene expression profile and normal lipid storage in their adipose tissue [149, 150]. These studies suggest that SREBP-1 is involved in the brown adipocyte lipogenesis and triglyceride storage when the excessive energy resources are available; however, SREBP-1's

In summary, these studies suggest that in activated brown adipocytes, glycogenesis and lipogenesis are upregulated to store/restore energy substrates, which is parallel with energy substrate metabolism and thermogenesis. These coordinated anabolic and catabolic processes are important to maintain the brown adipocyte

**5. The proceeding of targeting BAT in human metabolic disorders**

The understating of BAT energy homeostasis and the discovery of the functional BAT in humans lead to significant interests in targeting BAT for metabolic disorders, for example, obesity, insulin resistance, type 2 diabetes, and lipid profile abnormality. The ability of BAT metabolizing fatty acid and glucose from the intracellular storage, the peripheral tissues liberation, and teh dietary nutrition absorption makes it a good potential therapeutic target for combating metabolic disorders in humans. In addition to the BAT, beige/brite fat, which is coexisted in white adipose tissue,

function can be compensated by other factors when it is absent.

#### *Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

*Cellular Metabolism and Related Disorders*

34% respectively, in brown adipocytes [6]. These studies suggested the importance of glucose metabolism in brown adipocyte function. However, another detailed study indicated that glucose oxidation does not play a major role in brown adipocyte metabolism and thermogenesis, by showing that the rate of 14CO2 formation from the 14C glucose was relatively small compared with the maximum rate of oxygen consumption in activated brown adipocytes [125]. Studies using radio-labeled glucose also suggested that glucose uptake was only sufficient to fuel maximally ~15% of the thermogenic capacity in activated rodent brown adipocytes, suggesting that the significantly upregulated glucose uptake in activated brown adipocytes is disassociated with the relative low glucose-mediated thermogenic capacity [123, 125, 126]. A more recent study indicates that the brown adipocyte energy production from glucose depends on the state of the cell: glucose and fatty acid contribute equally to brown adipocyte oxygen consumption under the basal condition; upon catecholamine-

activation, oxygen consumption is mainly fueled by fatty acids [6].

The dissociation between high glucose uptake and low glucose-mediated thermogenesis in activated BAT raises an important question: what is the function of the intracellular glucose in brown adipocytes? The importance of glucose in the *de novo* lipogenesis in brown adipocytes has been reported [127–129]. Studies indicate that glucose uptake is an independent process of thermogenesis in both cold-challenged and catecholamine-activated BAT, which further supports that glucose uptake might play other role as energy substrate in activated brown adipocytes [116, 117, 130–132]. One detailed *in vitro* study using rat brown adipocytes showed that norepinephrine significantly enhanced glucose oxidation by sevenfold, while it also inhibited lipogenesis at the same time. On the other hand, insulin stimulation increased the lipogenesis by sevenfold in brown adipocytes whereas glucose oxidation remained very low. Most interestingly, the addition of insulin to the norepinephrine only potentiated the enhanced glucose oxidation, but do not enhance the lipogenesis [115]. On the other hand, another study showed that brown adipocytes converted a greater proportion of metabolized glucose into lactate and pyruvate, but only a smaller proportion into fatty acids through insulin-mediated pathway [125]. These studies suggest that glucose metabolism is involved in two different states in brown adipocytes, the thermogenic state and nonthermogenic state. It is plausible that during the thermogenic state, glucose contributes to both the energy production and lipogenesis; and during the nonthermogenic state, glucose contributes mainly to the lipogenesis and energy storage process in brown adipocytes, which explains the disassociation of glucose uptake and glucose metabolism in brown adipocytes. In addition to glucose, other energy substrates, including glycerol and amino acid (glutamate), might also contribute to BAT metabolism and thermogenesis in human and rodent brown adipocytes. However, the relative contribution and partition of these energy substrates are unclear [132–135].

The cold challenge not only enhances catabolic processes mediating energy substrate metabolism and heat generation, but also induces anabolic processes mediating fatty acid synthesis and lipogenesis, as well as glycogenesis [36, 111, 136–139]. Glucose can be stored as glycogen in brown adipocytes [36, 139]. Studies showed

that glycogen synthases (GStot) and uridine diphosphate glucose pyrophosphorylase (Udgp), which mediates glycogenesis, were upregulated upon the cold challenge in BAT [36, 139]. Interestingly, glycogen hydrolysis enzyme, glycogen phosphorylase (Pygl), was also upregulated after cold challenge [36]. Although cold-challenge upregulates both glycogen synthesis and degradation, it is reported

**16**

**4. Energy storage in BAT**

that the stored glycogen is used up shortly after the cold challenge [139]. Indicating that glycogen is not an efficient format for energy storage and a sustainable energy resource in brown adipocytes.

The glucose uptaken by brown adipocyte can also be converted to fatty acid through the *de novo* lipogenesis. Carbohydrate response element-binding protein (ChREBP) is the major transcription factor mediating fatty acid synthesis in adipocytes. There are two isoforms of ChREBP, ChREBPα and ChREBPβ. ChREBPα is abundantly expressed in BAT from rodents and humans [137, 140, 141]. *De novo* lipogenesis involves a series of enzymes mediating the sequential reactions converting glucose-derived citrate into fatty acids, including ATP-citrate lyase (ACLY), acetyl-CoA carboxylases 1 (ACC1), fatty acid synthase (FASN), and stearoyl-CoA desaturase-1 (SCD1). ChREBPα, coordinates with another transcription factor Max-like protein (MLX), directly binds to the carbohydrate response elements (ChoRE) and upregulates these *de novo* lipogenic genes expressions. ChREBPα can also increase the expression of ChREBPβ to further activate the *de novo* lipogenesis enzymes [142, 143]. It has been reported that lipogenic genes and AKT2-ChREBP pathways are upregulated to optimize fuel storage and thermogenesis upon cold stimulation in BAT [136]. In accordance with these studies, *ChREBP−/−* mice presented less BAT weight [142], and adipose-specific *ChREBP-knockout* mice had decreased carbohydrate-induced lipogenesis in BAT [144]. These studies indicate that ChREBP plays important roles in brown adipocyte's *de novo* lipogenesis and energy storage.

Sterol regulatory element-binding protein-1 (SREBP-1) is another transcription factor mediating the *de novo* lipogenesis [145]. Of the three different SREBP isoforms, SREBP1c is more abundant and SREBP1a is less abundant in the adipose tissue [137, 145, 146]. *In vitro* study showed that SREBP1c is sufficient to regulate lipogenic enzymes in cultured adipocytes [147]. In the adipose-specific *ap2- SREBP1c* transgenic mice, lipogenic enzyme Acc1, FASN and Scd1 expression as well as fatty acids synthesis rate were significantly upregulated in brown adipocytes [148]. In line with this study, *ap2-SREBP1a* transgenic mice also developed adipose tissue hypertrophy in accordance with an increased lipogenic enzyme profile and enhance *de novo* lipogenesis in the BAT [148]. However, some *in vivo* studies indicated that SREBP1c's role in the *de novo* lipogenesis in adipocytes is dispensable, as evidenced by *SREBP1-knockout* mice have normal lipogenic enzymes gene expression profile and normal lipid storage in their adipose tissue [149, 150]. These studies suggest that SREBP-1 is involved in the brown adipocyte lipogenesis and triglyceride storage when the excessive energy resources are available; however, SREBP-1's function can be compensated by other factors when it is absent.

In summary, these studies suggest that in activated brown adipocytes, glycogenesis and lipogenesis are upregulated to store/restore energy substrates, which is parallel with energy substrate metabolism and thermogenesis. These coordinated anabolic and catabolic processes are important to maintain the brown adipocyte energy homeostasis.

#### **5. The proceeding of targeting BAT in human metabolic disorders**

The understating of BAT energy homeostasis and the discovery of the functional BAT in humans lead to significant interests in targeting BAT for metabolic disorders, for example, obesity, insulin resistance, type 2 diabetes, and lipid profile abnormality. The ability of BAT metabolizing fatty acid and glucose from the intracellular storage, the peripheral tissues liberation, and teh dietary nutrition absorption makes it a good potential therapeutic target for combating metabolic disorders in humans. In addition to the BAT, beige/brite fat, which is coexisted in white adipose tissue,

can be recruited and activated (browning) in respond to cold challenge or pharmacological stimulation and serves as a target for metabolic disorders [151–154].

It has been reported that the BAT 18F-FDG uptake in humans correlates inversely with aging, adiposity, diabetic status, and BMI, indicating that the manipulation of BAT function is a possible approach for combating metabolic disorders [8–14, 16, 21–24, 155, 156]. The studies in mouse models and humans provide evidences for the metabolic benefit of BAT. Mice with genetic ablation of BAT and the *UCP1-knockout* mice under thermoneutrality, both developed obesity [18–20, 85]. In addition, BAT activation reduced hypercholesterolemia and protected mice from atherosclerosis development and liver steatosis [18–20, 85]. It is well-recognized that BAT can be activated upon seasonal temperature changes or short time period (several hours) mild cold challenge (16-19°C) in humans [7, 9, 10, 12, 21, 22, 24–27, 157]. However, the significance of BAT's contribution in human metabolism has not been clearly elucidated. Human studies indicated that BAT activity is inversely correlated with body fat deposition, suggests BAT can serve as a target for obesity [9, 10, 16, 21–24]. Studies also showed that fasting glucose was lower in the human subjects with higher BAT prevalence and the BAT activity were blunted in subjects with obesity [11, 16, 155]. More interestingly, in the same patients with multiple PET scans, BAT was more detectable when fasting glucose in the subjects were lower [16]. Other studies showed that a mild cold challenge significantly increased whole body glucose disposal, glucose oxidation, insulin sensitivity, and whole body energy expenditure in human subjects [13, 17, 158, 159]. Additional study showed that moderate cold challenge (18.06°C) significantly improved the peripheral glucose uptake and insulin sensitivity by 20%, but did not impact the pancreatic insulin secretion [17]. These studies strongly indicate the therapeutic potentials of targeting BAT for glucose metabolism in humans, but leave the question of the relative contribution of activated BAT in whole body glucose metabolism to be answered.

Cold challenge can also enhance BAT lipid metabolism. Studies showed that cold challenge led to significantly enhanced lipid mobilization, increased plasma fatty acid levels, as well as upregulated genes for lipid metabolism in human BAT [12, 58, 156, 160]. It is reported that circulating fatty acids were uptaken by BAT in cold-challenged humans by using the 18FTHA tracing method [88]. In addition, it has been shown that cold-activated BAT significantly contributed to whole body fatty acid utilization in healthy humans [17]. The fatty acid uptake is significantly lower in overweight human subjects compare to healthy humans [156]. Importantly, BAT activation upon mild cold challenge significantly increased systemic lipid metabolism, whole body lipolysis, triglyceride-fatty acid cycling, and fatty acid oxidation in overweight/obese subjects [14]. These studies suggest that BAT contributes to whole body lipid metabolism and homeostasis in healthy humans as well as in humans with metabolic disorders, suggesting that BAT can serve as a target for lipid abnormality in humans.

The significance of BAT/beige adipose tissue in human whole body metabolism has been studied and remarkable progress has been made in recent years. The acknowledgment that BAT can be activated and subsequently contributes to the human whole body energy expenditure is encouraging. Although the capacity and relative significance of BAT's contribution to whole body energy substrate metabolism has not been elucidated, it should be noticed that majority of the studies in humans were conducted for relative short time period with mild cold challenge conditions. Given, humans usually live under thermoneutrality, we can assume that their BAT functions are repressed under this condition. In addition, the prevalence of BAT in humans varies significantly, which depends on individual's life style, physical activity, and health conditions, which makes it harder to evaluate the contribution of BAT in whole body metabolism [7, 9–12, 21, 22, 24–27, 88, 156, 157].

**19**

provided the original work is properly cited.

\*Address all correspondence to: yuan.lu@case.edu

*Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

ders through BAT activation in the future.

metabolic disorder management.

**Acknowledgements**

to Mukesh K. Jain.

**Author details**

Center, Cleveland, Ohio, USA

Yuan Lu1,2\*

**Conflict of interest**

The author has no conflict of interest to declare.

**6. Conclusion**

Hence, a sustainable chronic mild cold challenge strategy aiming to recruit more BAT/beige adipose tissue and enhance their oxidative capacity might provide more significant therapeutic potential in humans over time, especially the human subjects with metabolic disorders. Future studies should also delineate the relative contribution from glucose and fatty acid in human BAT under physiological and pathological conditions; as well as compare the glucose and fatty acid partitioning in different tissues and organs, including skeletal muscle, heart, liver and BAT. These details will guide us to establish better strategies targeting metabolic disor-

The energy metabolism plays critical role in maintaining BAT thermogenic function in mammals. Through the energy metabolism, the chemical energy stored in nutrients (i.e., fatty acids and glucose) can be converted to thermoenergy and dissipated as heat in the BAT. Activated brown adipocytes not only contribute to the intracellular substrate homeostasis, but also contribute significantly to the whole body energy metabolism. BAT with functional thermoactivity is present in adult humans. Thus, BAT activation has remarkable therapeutic implications in human

This work was supported by American Heart Association National Scientist Development grant, AHA SDG grant, 12050558 to Yuan Lu; and NIH R35HL135789

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium,

1 Case Western Reserve University School of Medicine, Cleveland, Ohio, USA

2 Harrington Heart and Vascular Institute, University Hospitals Cleveland Medical

*Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

Hence, a sustainable chronic mild cold challenge strategy aiming to recruit more BAT/beige adipose tissue and enhance their oxidative capacity might provide more significant therapeutic potential in humans over time, especially the human subjects with metabolic disorders. Future studies should also delineate the relative contribution from glucose and fatty acid in human BAT under physiological and pathological conditions; as well as compare the glucose and fatty acid partitioning in different tissues and organs, including skeletal muscle, heart, liver and BAT. These details will guide us to establish better strategies targeting metabolic disorders through BAT activation in the future.

#### **6. Conclusion**

*Cellular Metabolism and Related Disorders*

can be recruited and activated (browning) in respond to cold challenge or pharmacological stimulation and serves as a target for metabolic disorders [151–154].

activated BAT in whole body glucose metabolism to be answered.

serve as a target for lipid abnormality in humans.

Cold challenge can also enhance BAT lipid metabolism. Studies showed that cold challenge led to significantly enhanced lipid mobilization, increased plasma fatty acid levels, as well as upregulated genes for lipid metabolism in human BAT [12, 58, 156, 160]. It is reported that circulating fatty acids were uptaken by BAT in cold-challenged humans by using the 18FTHA tracing method [88]. In addition, it has been shown that cold-activated BAT significantly contributed to whole body fatty acid utilization in healthy humans [17]. The fatty acid uptake is significantly lower in overweight human subjects compare to healthy humans [156]. Importantly, BAT activation upon mild cold challenge significantly increased systemic lipid metabolism, whole body lipolysis, triglyceride-fatty acid cycling, and fatty acid oxidation in overweight/obese subjects [14]. These studies suggest that BAT contributes to whole body lipid metabolism and homeostasis in healthy humans as well as in humans with metabolic disorders, suggesting that BAT can

The significance of BAT/beige adipose tissue in human whole body metabolism

has been studied and remarkable progress has been made in recent years. The acknowledgment that BAT can be activated and subsequently contributes to the human whole body energy expenditure is encouraging. Although the capacity and relative significance of BAT's contribution to whole body energy substrate metabolism has not been elucidated, it should be noticed that majority of the studies in humans were conducted for relative short time period with mild cold challenge conditions. Given, humans usually live under thermoneutrality, we can assume that their BAT functions are repressed under this condition. In addition, the prevalence of BAT in humans varies significantly, which depends on individual's life style, physical activity, and health conditions, which makes it harder to evaluate the contribution of BAT in whole body metabolism [7, 9–12, 21, 22, 24–27, 88, 156, 157].

It has been reported that the BAT 18F-FDG uptake in humans correlates inversely with aging, adiposity, diabetic status, and BMI, indicating that the manipulation of BAT function is a possible approach for combating metabolic disorders [8–14, 16, 21–24, 155, 156]. The studies in mouse models and humans provide evidences for the metabolic benefit of BAT. Mice with genetic ablation of BAT and the *UCP1-knockout* mice under thermoneutrality, both developed obesity [18–20, 85]. In addition, BAT activation reduced hypercholesterolemia and protected mice from atherosclerosis development and liver steatosis [18–20, 85]. It is well-recognized that BAT can be activated upon seasonal temperature changes or short time period (several hours) mild cold challenge (16-19°C) in humans [7, 9, 10, 12, 21, 22, 24–27, 157]. However, the significance of BAT's contribution in human metabolism has not been clearly elucidated. Human studies indicated that BAT activity is inversely correlated with body fat deposition, suggests BAT can serve as a target for obesity [9, 10, 16, 21–24]. Studies also showed that fasting glucose was lower in the human subjects with higher BAT prevalence and the BAT activity were blunted in subjects with obesity [11, 16, 155]. More interestingly, in the same patients with multiple PET scans, BAT was more detectable when fasting glucose in the subjects were lower [16]. Other studies showed that a mild cold challenge significantly increased whole body glucose disposal, glucose oxidation, insulin sensitivity, and whole body energy expenditure in human subjects [13, 17, 158, 159]. Additional study showed that moderate cold challenge (18.06°C) significantly improved the peripheral glucose uptake and insulin sensitivity by 20%, but did not impact the pancreatic insulin secretion [17]. These studies strongly indicate the therapeutic potentials of targeting BAT for glucose metabolism in humans, but leave the question of the relative contribution of

**18**

The energy metabolism plays critical role in maintaining BAT thermogenic function in mammals. Through the energy metabolism, the chemical energy stored in nutrients (i.e., fatty acids and glucose) can be converted to thermoenergy and dissipated as heat in the BAT. Activated brown adipocytes not only contribute to the intracellular substrate homeostasis, but also contribute significantly to the whole body energy metabolism. BAT with functional thermoactivity is present in adult humans. Thus, BAT activation has remarkable therapeutic implications in human metabolic disorder management.

#### **Acknowledgements**

This work was supported by American Heart Association National Scientist Development grant, AHA SDG grant, 12050558 to Yuan Lu; and NIH R35HL135789 to Mukesh K. Jain.

#### **Conflict of interest**

The author has no conflict of interest to declare.

#### **Author details**

Yuan Lu1,2\*

1 Case Western Reserve University School of Medicine, Cleveland, Ohio, USA

2 Harrington Heart and Vascular Institute, University Hospitals Cleveland Medical Center, Cleveland, Ohio, USA

\*Address all correspondence to: yuan.lu@case.edu

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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**20**

*Cellular Metabolism and Related Disorders*

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2007;**5**(4):279-291

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[102] Harmon CM, Abumrad NA. Binding of sulfosuccinimidyl fatty acids to adipocyte membrane proteins: Isolation and amino-terminal sequence of an 88-kD protein implicated in transport of long-chain fatty acids. The Journal of Membrane Biology.

[103] Tao N, Wagner SJ, Lublin DM. CD36 is palmitoylated on both

1993;**133**(1):43-49

*Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

of triglyceride-rich lipoproteins by interaction with proteoglycanbound lipoprotein lipase. The Journal of Biological Chemistry. 2005;**280**(22):21553-21560

*Cellular Metabolism and Related Disorders*

heart: Isolation and characterization of its cDNA clone. FEBS Letters.

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[79] Gonzalez-Hurtado E et al. Fatty acid oxidation is required for active and quiescent brown adipose tissue maintenance and thermogenic programing. Molecular Metabolism.

[80] Schuler AM et al. Synergistic heterozygosity in mice with inherited enzyme deficiencies of

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[81] Thorpe C, Kim JJ. Structure and mechanism of action of the acyl-CoA dehydrogenases. The FASEB Journal.

[82] Gregersen N et al. Mutation analysis in mitochondrial fatty acid oxidation defects: Exemplified by acyl-CoA dehydrogenase deficiencies, with special focus on genotype-phenotype relationship. Human Mutation.

[83] Bartelt A et al. Brown adipose tissue activity controls triglyceride clearance. Nature Medicine. 2011;**17**(2):200-205

[84] Khedoe PP et al. Brown adipose tissue takes up plasma triglycerides mostly after lipolysis. Journal of Lipid

[85] Berbee JF et al. Brown fat activation

Research. 2015;**56**(1):51-59

2015;**6**:6356

reduces hypercholesterolaemia and protects from atherosclerosis development. Nature Communications.

[86] Merkel M et al. Apolipoprotein AV accelerates plasma hydrolysis

1995;**363**(1-2):41-45

2018;**7**:45-56

2005;**85**(1):7-11

1995;**9**(9):718-725

2001;**18**(3):169-189

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[74] Syamsunarno MR et al. Fatty acid binding protein 4 and 5 play a crucial role in thermogenesis under the conditions of fasting and cold stress.

PLoS One. 2014;**9**(6):e90825

1996;**271**(12):6972-6977

[77] Yamazaki N et al. High expression of a novel carnitine palmitoyltransferase I like protein in rat brown adipose tissue and

[76] Nyman LR et al. Homozygous carnitine palmitoyltransferase 1a (liver isoform) deficiency is lethal in the mouse. Molecular Genetics and Metabolism. 2005;**86**(1-2):179-187

[75] Esser V et al. Expression of a cDNA isolated from rat brown adipose tissue and heart identifies the product as the muscle isoform of carnitine palmitoyltransferase I (M-CPT I). M-CPT I is the predominant CPT I isoform expressed in both white (epididymal) and brown adipocytes. The Journal of Biological Chemistry.

1999;**96**(10):5528-5532

1999;**40**(5):967-972

[70] Makowski L, Hotamisligil GS. The role of fatty acid binding proteins in metabolic syndrome and atherosclerosis. Current Opinion in Lipidology. 2005;**16**(5):543-548

**24**

[87] Pennacchio LA et al. An apolipoprotein influencing triglycerides in humans and mice revealed by comparative sequencing. Science. 2001;**294**(5540):169-173

[88] Blondin DP et al. Dietary fatty acid metabolism of brown adipose tissue in cold-acclimated men. Nature Communications. 2017;**8**:14146

[89] Amri EZ et al. Maturation and secretion of lipoprotein lipase in cultured adipose cells. II. Effects of tunicamycin on activation and secretion of the enzyme. Biochimica et Biophysica Acta. 1986;**875**(2):334-343

[90] Carneheim C, Nedergaard J, Cannon B. Cold-induced betaadrenergic recruitment of lipoprotein lipase in brown fat is due to increased transcription. The American Journal of Physiology. 1988;**254**(2 Pt 1):E155-E161

[91] Ong JM, Kern PA. The role of glucose and glycosylation in the regulation of lipoprotein lipase synthesis and secretion in rat adipocytes. The Journal of Biological Chemistry. 1989;**264**(6):3177-3182

[92] Stahl A. A current review of fatty acid transport proteins (SLC27). Pflügers Archiv. 2004;**447**(5):722-727

[93] Anderson CM, Stahl A. SLC27 fatty acid transport proteins. Molecular Aspects of Medicine. 2013;**34**(2-3):516-528

[94] Wang H, Eckel RH. Lipoprotein lipase: From gene to obesity. American Journal of Physiology. Endocrinology and Metabolism. 2009;**297**(2):E271-E288

[95] Beigneux AP et al. Glycosylphosphatidylinositolanchored high-density lipoproteinbinding protein 1 plays a critical role in the lipolytic processing of chylomicrons. Cell Metabolism. 2007;**5**(4):279-291

[96] Davies BS et al. GPIHBP1 is responsible for the entry of lipoprotein lipase into capillaries. Cell Metabolism. 2010;**12**(1):42-52

[97] Beisiegel U, Weber W, Bengtsson-Olivecrona G. Lipoprotein lipase enhances the binding of chylomicrons to low density lipoprotein receptorrelated protein. Proceedings of the National Academy of Sciences of the United States of America. 1991;**88**(19):8342-8346

[98] Mead JR, Irvine SA, Ramji DP. Lipoprotein lipase: Structure, function, regulation, and role in disease. Journal of Molecular Medicine (Berlin, Germany). 2002;**80**(12):753-769

[99] Wang CS, Hartsuck J, McConathy WJ. Structure and functional properties of lipoprotein lipase. Biochimica et Biophysica Acta. 1992;**1123**(1):1-17

[100] Wu Q et al. Fatty acid transport protein 1 is required for nonshivering thermogenesis in brown adipose tissue. Diabetes. 2006;**55**(12):3229-3237

[101] Wu Q et al. FATP1 is an insulinsensitive fatty acid transporter involved in diet-induced obesity. Molecular and Cellular Biology. 2006;**26**(9):3455-3467

[102] Harmon CM, Abumrad NA. Binding of sulfosuccinimidyl fatty acids to adipocyte membrane proteins: Isolation and amino-terminal sequence of an 88-kD protein implicated in transport of long-chain fatty acids. The Journal of Membrane Biology. 1993;**133**(1):43-49

[103] Tao N, Wagner SJ, Lublin DM. CD36 is palmitoylated on both N- and C-terminal cytoplasmic tails. The Journal of Biological Chemistry. 1996;**271**(37):22315-22320

[104] Anderson CM et al. Dependence of brown adipose tissue function on CD36-mediated coenzyme Q uptake. Cell Reports. 2015;**10**(4):505-515

[105] Som P et al. A fluorinated glucose analog, 2-fluoro-2-deoxy-D-glucose (F-18): Nontoxic tracer for rapid tumor detection. Journal of Nuclear Medicine. 1980;**21**(7):670-675

[106] Kelloff GJ et al. Progress and promise of FDG-PET imaging for cancer patient management and oncologic drug development. Clinical Cancer Research. 2005;**11**(8):2785-2808

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[108] Vallerand AL, Perusse F, Bukowiecki LJ. Cold exposure potentiates the effect of insulin on in vivo glucose uptake. The American Journal of Physiology. 1987;**253**(2 Pt 1):E179-E186

[109] Vallerand AL, Perusse F, Bukowiecki LJ. Stimulatory effects of cold exposure and cold acclimation on glucose uptake in rat peripheral tissues. The American Journal of Physiology. 1990;**259**(5 Pt 2):R1043-R1049

[110] Shimizu Y, Nikami H, Saito M. Sympathetic activation of glucose utilization in brown adipose tissue in rats. Journal of Biochemistry. 1991;**110**(5):688-692

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from differential gene expression and confirmation in vivo. The FASEB Journal. 2002;**16**(2):155-168

[112] Shimizu Y et al. Increased expression of glucose transporter GLUT-4 in brown adipose tissue of fasted rats after cold exposure. The American Journal of Physiology. 1993;**264**(6 Pt 1):E890-E895

[113] Wang X, Wahl R. Responses of the insulin signaling pathways in the brown adipose tissue of rats following cold exposure. PLoS One. 2014;**9**(6):e99772

[114] Shibata H et al. Cold exposure reverses inhibitory effects of fasting on peripheral glucose uptake in rats. The American Journal of Physiology. 1989;**257**(1 Pt 2):R96-R101

[115] Ebner S et al. Effects of insulin and norepinephrine on glucose transport and metabolism in rat brown adipocytes. Potentiation by insulin of norepinephrine-induced glucose oxidation. European Journal of Biochemistry. 1987;**170**(1-2):469-474

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[117] Olsen JM et al. Glucose uptake in brown fat cells is dependent on mTOR complex 2-promoted GLUT1 translocation. The Journal of Cell Biology. 2014;**207**(3):365-374

[118] Bell GI et al. Molecular biology of mammalian glucose transporters. Diabetes Care. 1990;**13**(3):198-208

[119] Gatenby RA, Gillies RJ. Why do cancers have high aerobic glycolysis? Nature Reviews. Cancer. 2004;**4**(11):891-899

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[123] Ma SW, Foster DO. Uptake of glucose and release of fatty acids and glycerol by rat brown adipose tissue in vivo. Canadian Journal of Physiology and Pharmacology. 1986;**64**(5):609-614

[124] Jeong JH, Chang JS, Jo YH.

Intracellular glycolysis in brown adipose tissue is essential for optogenetically induced nonshivering thermogenesis in mice. Scientific Reports. 2018;**8**(1):6672

[125] Saggerson ED, McAllister TW, Baht HS. Lipogenesis in rat brown adipocytes. Effects of insulin and noradrenaline, contributions from glucose and lactate as precursors and comparisons with white adipocytes. The Biochemical Journal. 1988;**251**(3):701-709

[126] Isler D, Hill HP, Meier MK. Glucose

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The Biochemical Journal. 1987;**245**(3):789-793

1992;**185**(3):1078-1082

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119-124

2):485-490

#### *Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

*Cellular Metabolism and Related Disorders*

N- and C-terminal cytoplasmic tails. The Journal of Biological Chemistry. from differential gene expression and confirmation in vivo. The FASEB

Journal. 2002;**16**(2):155-168

[112] Shimizu Y et al. Increased expression of glucose transporter GLUT-4 in brown adipose tissue of fasted rats after cold exposure. The American Journal of Physiology. 1993;**264**(6 Pt 1):E890-E895

[113] Wang X, Wahl R. Responses of the insulin signaling pathways in the brown adipose tissue of rats following cold exposure. PLoS One. 2014;**9**(6):e99772

[114] Shibata H et al. Cold exposure reverses inhibitory effects of fasting on peripheral glucose uptake in rats. The American Journal of Physiology.

[115] Ebner S et al. Effects of insulin and norepinephrine on glucose transport and metabolism in rat brown adipocytes. Potentiation by insulin of norepinephrine-induced glucose oxidation. European Journal of Biochemistry. 1987;**170**(1-2):469-474

[116] Hutchinson DS et al. Betaadrenoceptors, but not alpha-

adrenoceptors, stimulate AMP-activated protein kinase in brown adipocytes independently of uncoupling protein-1. Diabetologia. 2005;**48**(11):2386-2395

[117] Olsen JM et al. Glucose uptake in brown fat cells is dependent on mTOR complex 2-promoted GLUT1 translocation. The Journal of Cell Biology. 2014;**207**(3):365-374

[118] Bell GI et al. Molecular biology of mammalian glucose transporters. Diabetes Care. 1990;**13**(3):198-208

[119] Gatenby RA, Gillies RJ. Why do cancers have high aerobic glycolysis? Nature Reviews. Cancer.

[120] Marette A, Bukowiecki LJ. Noradrenaline stimulates glucose

2004;**4**(11):891-899

1989;**257**(1 Pt 2):R96-R101

[104] Anderson CM et al. Dependence of brown adipose tissue function on CD36-mediated coenzyme Q uptake. Cell Reports. 2015;**10**(4):505-515

[105] Som P et al. A fluorinated glucose analog, 2-fluoro-2-deoxy-D-glucose (F-18): Nontoxic tracer for rapid tumor detection. Journal of Nuclear Medicine.

[106] Kelloff GJ et al. Progress and promise of FDG-PET imaging for cancer patient management and oncologic drug development. Clinical Cancer Research.

[107] Greco-Perotto R et al. Stimulatory effect of cold adaptation on glucose utilization by brown adipose tissue. Relationship with changes in the glucose transporter system. The Journal of Biological Chemistry.

1996;**271**(37):22315-22320

1980;**21**(7):670-675

2005;**11**(8):2785-2808

1987;**262**(16):7732-7736

1):E179-E186

[108] Vallerand AL, Perusse F, Bukowiecki LJ. Cold exposure potentiates the effect of insulin on in vivo glucose uptake. The American Journal of Physiology. 1987;**253**(2 Pt

[109] Vallerand AL, Perusse F,

Bukowiecki LJ. Stimulatory effects of cold exposure and cold acclimation on glucose uptake in rat peripheral tissues. The American Journal of Physiology. 1990;**259**(5 Pt 2):R1043-R1049

[110] Shimizu Y, Nikami H, Saito M. Sympathetic activation of glucose utilization in brown adipose tissue in rats. Journal of Biochemistry.

[111] Yu XX et al. Cold elicits the simultaneous induction of fatty acid synthesis and beta-oxidation in murine brown adipose tissue: Prediction

1991;**110**(5):688-692

**26**

transport in rat brown adipocytes by activating thermogenesis. Evidence that fatty acid activation of mitochondrial respiration enhances glucose transport. The Biochemical Journal. 1991;**277**(Pt 1): 119-124

[121] Lee P et al. Brown adipose tissue exhibits a glucose-responsive thermogenic biorhythm in humans. Cell Metabolism. 2016;**23**(4):602-609

[122] Shimizu Y et al. Noradrenaline increases glucose transport into brown adipocytes in culture by a mechanism different from that of insulin. The Biochemical Journal. 1996;**314**(Pt 2):485-490

[123] Ma SW, Foster DO. Uptake of glucose and release of fatty acids and glycerol by rat brown adipose tissue in vivo. Canadian Journal of Physiology and Pharmacology. 1986;**64**(5):609-614

[124] Jeong JH, Chang JS, Jo YH. Intracellular glycolysis in brown adipose tissue is essential for optogenetically induced nonshivering thermogenesis in mice. Scientific Reports. 2018;**8**(1):6672

[125] Saggerson ED, McAllister TW, Baht HS. Lipogenesis in rat brown adipocytes. Effects of insulin and noradrenaline, contributions from glucose and lactate as precursors and comparisons with white adipocytes. The Biochemical Journal. 1988;**251**(3):701-709

[126] Isler D, Hill HP, Meier MK. Glucose metabolism in isolated brown adipocytes under beta-adrenergic stimulation. Quantitative contribution of glucose to total thermogenesis. The Biochemical Journal. 1987;**245**(3):789-793

[127] Nikami H et al. Cold exposure increases glucose utilization and glucose transporter expression in brown adipose tissue. Biochemical and Biophysical Research Communications. 1992;**185**(3):1078-1082

[128] Smith DM et al. Glucose transporter expression and glucose utilization in skeletal muscle and brown adipose tissue during starvation and re-feeding. The Biochemical Journal. 1992;**282**(Pt 1):231-235

[129] Sugden MC, Holness MJ. Physiological modulation of the uptake and fate of glucose in brown adipose tissue. The Biochemical Journal. 1993;**295**(Pt 1):171-176

[130] Hankir MK et al. Dissociation between brown adipose tissue (18) F-FDG uptake and thermogenesis in uncoupling protein 1-deficient mice. Journal of Nuclear Medicine. 2017;**58**(7):1100-1103

[131] Olsen JM et al. beta3- Adrenergically induced glucose uptake in brown adipose tissue is independent of UCP1 presence or activity: Mediation through the mTOR pathway. Molecular Metabolism. 2017;**6**(6):611-619

[132] Weir G et al. Substantial metabolic activity of human brown adipose tissue during warm conditions and coldinduced lipolysis of local triglycerides. Cell Metabolism. 2018;**27**(6):1348 e4-1355 e4

[133] Chakrabarty K, Chaudhuri B, Jeffay H. Glycerokinase activity in human brown adipose tissue. Journal of Lipid Research. 1983;**24**(4):381-390

[134] Kawashita NH et al. Glycerokinase activity in brown adipose tissue: A sympathetic regulation? American Journal of Physiology. Regulatory, Integrative and Comparative Physiology. 2002;**282**(4):R1185-R1190

[135] Festuccia WT et al. Control of glyceroneogenic activity in rat brown adipose tissue. American Journal of Physiology. Regulatory, Integrative and Comparative Physiology. 2003;**285**(1):R177-R182

[136] Sanchez-Gurmaches J et al. Brown fat AKT2 is a cold-induced kinase that stimulates ChREBP-mediated *De novo* lipogenesis to optimize fuel storage and thermogenesis. Cell Metabolism. 2018;**27**(1):195 e6-209 e6

[137] Song Z, Xiaoli AM, Yang F. Regulation and metabolic significance of *De novo* lipogenesis in adipose tissues. Nutrients. 2018;**10**(10):1383

[138] Guilherme A et al. Adipocyte lipid synthesis coupled to neuronal control of thermogenic programming. Molecular Metabolism. 2017;**6**(8):781-796

[139] Farkas V, Kelenyi G, Sandor A. A dramatic accumulation of glycogen in the brown adipose tissue of rats following recovery from cold exposure. Archives of Biochemistry and Biophysics. 1999;**365**(1):54-61

[140] Hurtado del Pozo C et al. ChREBP expression in the liver, adipose tissue and differentiated preadipocytes in human obesity. Biochimica et Biophysica Acta. 2011;**1811**(12):1194-1200

[141] Witte N et al. The glucose sensor ChREBP links *De novo* lipogenesis to PPARgamma activity and adipocyte differentiation. Endocrinology. 2015;**156**(11):4008-4019

[142] Iizuka K et al. Deficiency of carbohydrate response element-binding protein (ChREBP) reduces lipogenesis as well as glycolysis. Proceedings of the National Academy of Sciences of the United States of America. 2004;**101**(19):7281-7286

[143] Herman MA et al. A novel ChREBP isoform in adipose tissue regulates systemic glucose metabolism. Nature. 2012;**484**(7394):333-338

[144] Vijayakumar A et al. Absence of carbohydrate response element binding protein in adipocytes causes systemic insulin resistance and impairs glucose transport. Cell Reports. 2017;**21**(4):1021-1035

[145] Eberle D et al. SREBP transcription factors: Master regulators of lipid homeostasis. Biochimie. 2004;**86**(11):839-848

[146] Osborne TF. Sterol regulatory element-binding proteins (SREBPs): Key regulators of nutritional homeostasis and insulin action. The Journal of Biological Chemistry. 2000;**275**(42):32379-32382

[147] Kim JB, Spiegelman BM. ADD1/ SREBP1 promotes adipocyte differentiation and gene expression linked to fatty acid metabolism. Genes & Development. 1996;**10**(9):1096-1107

[148] Horton JD et al. Overexpression of sterol regulatory element-binding protein-1a in mouse adipose tissue produces adipocyte hypertrophy, increased fatty acid secretion, and fatty liver. The Journal of Biological Chemistry. 2003;**278**(38):36652-36660

[149] Shimano H et al. Elevated levels of SREBP-2 and cholesterol synthesis in livers of mice homozygous for a targeted disruption of the SREBP-1 gene. The Journal of Clinical Investigation. 1997;**100**(8):2115-2124

[150] Yahagi N et al. Absence of sterol regulatory element-binding protein-1 (SREBP-1) ameliorates fatty livers but not obesity or insulin resistance in Lep(ob)/Lep(ob) mice. The Journal of Biological Chemistry. 2002;**277**(22):19353-19357

[151] Guerra C et al. Emergence of brown adipocytes in white fat in mice is under genetic control. Effects on body weight and adiposity. The Journal of Clinical Investigation. 1998;**102**(2):412-420

[152] Cederberg A et al. FOXC2 is a winged helix gene that counteracts obesity, hypertriglyceridemia, and

**29**

*Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

diet-induced insulin resistance. Cell.

[153] Kiefer FW et al. Retinaldehyde

[154] Schulz TJ et al. Brown-fat paucity due to impaired BMP signalling induces compensatory browning of white fat. Nature. 2013;**495**(7441):379-383

[155] Orava J et al. Blunted metabolic

stimulation in brown adipose tissue of obese humans. Obesity (Silver Spring).

responses to cold and insulin

[156] Blondin DP et al. Selective impairment of glucose but not fatty acid or oxidative metabolism in brown adipose tissue of subjects with type 2 diabetes. Diabetes.

[157] Au-Yong IT et al. Brown adipose tissue and seasonal variation in humans.

Diabetes. 2009;**58**(11):2583-2587

[158] Leitner BP et al. Mapping of human brown adipose tissue in lean and obese young men. Proceedings of the National Academy of Sciences of the United States of America.

[159] Hanssen MJ et al. Short-term cold acclimation improves insulin sensitivity in patients with type 2 diabetes mellitus. Nature Medicine. 2015;**21**(8):863-865

[160] Hoeke G et al. Short-term cooling increases serum triglycerides and small high-density lipoprotein levels in humans. Journal of Clinical Lipidology.

2013;**21**(11):2279-2287

2015;**64**(7):2388-2397

2017;**114**(32):8649-8654

2017;**11**(4):920 e2-928 e2

dehydrogenase 1 regulates a thermogenic program in white adipose tissue. Nature Medicine.

2001;**106**(5):563-573

2012;**18**(6):918-925

*Brown Adipose Tissue Energy Metabolism DOI: http://dx.doi.org/10.5772/intechopen.83712*

diet-induced insulin resistance. Cell. 2001;**106**(5):563-573

*Cellular Metabolism and Related Disorders*

2018;**27**(1):195 e6-209 e6

[137] Song Z, Xiaoli AM, Yang F. Regulation and metabolic significance of *De novo* lipogenesis in adipose tissues.

[138] Guilherme A et al. Adipocyte lipid synthesis coupled to neuronal control of thermogenic programming. Molecular

Nutrients. 2018;**10**(10):1383

Metabolism. 2017;**6**(8):781-796

Archives of Biochemistry and Biophysics. 1999;**365**(1):54-61

Acta. 2011;**1811**(12):1194-1200

[142] Iizuka K et al. Deficiency of carbohydrate response element-binding protein (ChREBP) reduces lipogenesis as well as glycolysis. Proceedings of the National Academy of Sciences of the United States of America.

[143] Herman MA et al. A novel ChREBP isoform in adipose tissue regulates systemic glucose metabolism. Nature.

[144] Vijayakumar A et al. Absence of carbohydrate response element binding protein in adipocytes causes systemic insulin resistance and impairs

2004;**101**(19):7281-7286

2012;**484**(7394):333-338

[139] Farkas V, Kelenyi G, Sandor A. A dramatic accumulation of glycogen in the brown adipose tissue of rats following recovery from cold exposure.

[140] Hurtado del Pozo C et al. ChREBP expression in the liver, adipose tissue and differentiated preadipocytes in human obesity. Biochimica et Biophysica

[141] Witte N et al. The glucose sensor ChREBP links *De novo* lipogenesis to PPARgamma activity and adipocyte differentiation. Endocrinology. 2015;**156**(11):4008-4019

[136] Sanchez-Gurmaches J et al. Brown fat AKT2 is a cold-induced kinase that stimulates ChREBP-mediated *De novo* lipogenesis to optimize fuel storage and thermogenesis. Cell Metabolism.

glucose transport. Cell Reports.

transcription factors: Master regulators of lipid homeostasis. Biochimie.

[146] Osborne TF. Sterol regulatory element-binding proteins (SREBPs):

[147] Kim JB, Spiegelman BM. ADD1/

[148] Horton JD et al. Overexpression of sterol regulatory element-binding protein-1a in mouse adipose tissue produces adipocyte hypertrophy, increased fatty acid secretion, and fatty liver. The Journal of Biological Chemistry. 2003;**278**(38):36652-36660

[149] Shimano H et al. Elevated levels of SREBP-2 and cholesterol synthesis in livers of mice homozygous for a targeted disruption of the SREBP-1 gene. The Journal of Clinical Investigation.

1997;**100**(8):2115-2124

[150] Yahagi N et al. Absence of sterol regulatory element-binding protein-1 (SREBP-1) ameliorates fatty livers but not obesity or insulin resistance in Lep(ob)/Lep(ob) mice. The Journal of Biological Chemistry.

2002;**277**(22):19353-19357

[151] Guerra C et al. Emergence of brown adipocytes in white fat in mice is under genetic control. Effects on body weight and adiposity. The Journal of Clinical Investigation. 1998;**102**(2):412-420

[152] Cederberg A et al. FOXC2 is a winged helix gene that counteracts obesity, hypertriglyceridemia, and

Key regulators of nutritional homeostasis and insulin action. The Journal of Biological Chemistry. 2000;**275**(42):32379-32382

SREBP1 promotes adipocyte differentiation and gene expression linked to fatty acid metabolism. Genes & Development. 1996;**10**(9):1096-1107

2017;**21**(4):1021-1035

2004;**86**(11):839-848

[145] Eberle D et al. SREBP

**28**

[153] Kiefer FW et al. Retinaldehyde dehydrogenase 1 regulates a thermogenic program in white adipose tissue. Nature Medicine. 2012;**18**(6):918-925

[154] Schulz TJ et al. Brown-fat paucity due to impaired BMP signalling induces compensatory browning of white fat. Nature. 2013;**495**(7441):379-383

[155] Orava J et al. Blunted metabolic responses to cold and insulin stimulation in brown adipose tissue of obese humans. Obesity (Silver Spring). 2013;**21**(11):2279-2287

[156] Blondin DP et al. Selective impairment of glucose but not fatty acid or oxidative metabolism in brown adipose tissue of subjects with type 2 diabetes. Diabetes. 2015;**64**(7):2388-2397

[157] Au-Yong IT et al. Brown adipose tissue and seasonal variation in humans. Diabetes. 2009;**58**(11):2583-2587

[158] Leitner BP et al. Mapping of human brown adipose tissue in lean and obese young men. Proceedings of the National Academy of Sciences of the United States of America. 2017;**114**(32):8649-8654

[159] Hanssen MJ et al. Short-term cold acclimation improves insulin sensitivity in patients with type 2 diabetes mellitus. Nature Medicine. 2015;**21**(8):863-865

[160] Hoeke G et al. Short-term cooling increases serum triglycerides and small high-density lipoprotein levels in humans. Journal of Clinical Lipidology. 2017;**11**(4):920 e2-928 e2

**31**

**Chapter 3**

Its Rate

*Avital Schurr*

rates of oxygen (CMRO2

standing of brain energy metabolism.

oxygen, paradigm shift, polarography

**1.1 The first eight decades (1900–1980)**

**Abstract**

Cerebral Energy Metabolism:

Measuring and Understanding

The study of brain energy metabolism has taken second place to that of muscle ever since the dawn of this field of research. Consequently, each new discovery made using muscle tissue that advanced our understanding of the biochemistry of energy metabolic processes was attempted to be duplicated in brain tissue. It was only when the brain's high energy needs were recognized that researchers realized its vulnerability to any mishap in its energy supplies and that this vulnerability may play a role in various brain disorders. Understanding of the mechanisms by which the brain deals with energy shortage is of utmost importance in shedding light on the fundamentals of brain disorders and their potential treatment. To achieve such understanding, accurate measurement of brain energy metabolic rates is necessary. This chapter summarizes the history of the current knowledge of the biochemical processes responsible for the production of adenosine triphosphate (ATP) in the brain. It briefly reviews the various techniques used to measure cerebral metabolic

measuring the cerebral metabolic rate of lactate (CMRlactate) to improve our under-

Most human cells produce adenosine triphosphate (ATP) via two, mostly interconnected biochemical pathways, glycolysis and mitochondrial oxidative phosphorylation. Erythrocytes (red blood cells, RBCs) produce their ATP via the glycolytic pathway alone, since they lack mitochondria. Under anaerobic conditions, the less efficient glycolytic pathway is the main source of ATP supply, since without oxygen the oxidative phosphorylation pathway cannot be maintained. Throughout the first half of the twentieth century the majority of the researchers in the field of energy metabolism made muscle their tissue of choice for the study of energy metabolism. While muscle was believed to require a great deal of energy to perform its work, the brain was assumed to be a low consumer of energy, as indicated by the following quote: "*the brain is not a seat of active combustion, and considering the very small increase in CO*<sup>2</sup> *in the torcular blood it seems to us very improbable that the temperature* 

**Keywords:** BOLD fMRI, cerebral metabolic rate, glucose, glycolysis, lactate,

**1. A short review of brain energy metabolism research**

) and glucose (CMRglucose), and elaborates on the potential of

#### **Chapter 3**

## Cerebral Energy Metabolism: Measuring and Understanding Its Rate

*Avital Schurr*

#### **Abstract**

The study of brain energy metabolism has taken second place to that of muscle ever since the dawn of this field of research. Consequently, each new discovery made using muscle tissue that advanced our understanding of the biochemistry of energy metabolic processes was attempted to be duplicated in brain tissue. It was only when the brain's high energy needs were recognized that researchers realized its vulnerability to any mishap in its energy supplies and that this vulnerability may play a role in various brain disorders. Understanding of the mechanisms by which the brain deals with energy shortage is of utmost importance in shedding light on the fundamentals of brain disorders and their potential treatment. To achieve such understanding, accurate measurement of brain energy metabolic rates is necessary. This chapter summarizes the history of the current knowledge of the biochemical processes responsible for the production of adenosine triphosphate (ATP) in the brain. It briefly reviews the various techniques used to measure cerebral metabolic rates of oxygen (CMRO2 ) and glucose (CMRglucose), and elaborates on the potential of measuring the cerebral metabolic rate of lactate (CMRlactate) to improve our understanding of brain energy metabolism.

**Keywords:** BOLD fMRI, cerebral metabolic rate, glucose, glycolysis, lactate, oxygen, paradigm shift, polarography

#### **1. A short review of brain energy metabolism research**

#### **1.1 The first eight decades (1900–1980)**

Most human cells produce adenosine triphosphate (ATP) via two, mostly interconnected biochemical pathways, glycolysis and mitochondrial oxidative phosphorylation. Erythrocytes (red blood cells, RBCs) produce their ATP via the glycolytic pathway alone, since they lack mitochondria. Under anaerobic conditions, the less efficient glycolytic pathway is the main source of ATP supply, since without oxygen the oxidative phosphorylation pathway cannot be maintained. Throughout the first half of the twentieth century the majority of the researchers in the field of energy metabolism made muscle their tissue of choice for the study of energy metabolism. While muscle was believed to require a great deal of energy to perform its work, the brain was assumed to be a low consumer of energy, as indicated by the following quote: "*the brain is not a seat of active combustion, and considering the very small increase in CO*<sup>2</sup> *in the torcular blood it seems to us very improbable that the temperature* 

*of the brain should be perceptibly greater than that of the blood*" [1]. It was Tashiro [2] who was the first to demonstrate that nerve produces CO2 and ammonia during its metabolism. By 1924, Warburg et al. [3] demonstrated that brain tissue is able to convert large amount of glucose to lactic acid, and in 1927 it was shown that nerve produces a measurable amount of heat, an amount that increases upon electric stimulation [4]. Moreover, this increase in heat production was shown to correlate with the amount of oxygen consumed. According to Holmes [5] the above finding was the necessary proof that nerve impulse is a chemical process. All the major discoveries that have led to the elucidation of the biochemical pathways of energy metabolism were made using the tissue of choice in the field i.e., muscle. Both the tricarboxylic acid cycle and the glycolytic pathways were introduced in 1937 and 1940, respectively (the reader is directed to the many detailed reviews on the topic that are available). While there is a general agreement among biochemists, physiologists and neuroscientists as to the accuracy of the mitochondrial tricarboxylic acid (TCA) cycle and the oxidative phosphorylation pathway, disagreements exist on the accuracy of the glycolytic pathway. Hence, glycolysis is the main focus of this chapter, since the original drawing of the pathway stands in conflict with various research findings of the past three decades (for more detailed reviews of this topic the reader is directed to [6, 7]) . Although some research on brain energy metabolism was performed during 1920s and 1930s, it was limited to very few laboratories. Among them, that of Eric G. Holmes and Barbara E. Holmes pioneered important research in the field. The duo, who later joined by C.A. Ashford, published a series of papers [8–14] to demonstrate brain tissue production of lactate from glucose (similar to muscle metabolism), the involvement of phosphates and glycogen in this metabolism and the ability of brain tissue to oxidize lactate. Unfortunately, the latter never received neither the praise nor the scrutiny it deserved. This point is expanded upon in the next section, although it must be emphasized here that if the importance of that discovery would have been recognized at the time, our understanding of brain energy metabolism would be significantly accelerated and advanced. Nevertheless, Holmes and Ashford interpreted their finding of lactate oxidation simply as a process by which the brain rids itself of a waste product, since lactate was believed until the mid 1980s and even beyond, to be just that, a useless end-product of carbohydrate metabolism. Consequently, the research by Holmes and Ashford quickly became obscure. It would have stayed this way if not for a literature search I carried out in 2005 working on an upcoming paper that dealt with the possible role of lactate as an oxidative brain energy substrate [15]. Spending several weeks in the basement of my university medical school library (at the time most of the old literature has not yet been digitized) was an experience akin to treasure hunt, an experience I still cherish today. Discovering Holmes and Ashford's papers in 75-year old, dust-covered, heavy volumes of the Biochemical Journal was almost as exciting as conducting our own research [16]. Three years prior to the publication of the latter on brain energy metabolism, Brooks published his controversial work on muscle energy metabolism [17]. His proposal that skeletal muscle both produces and consumes lactate met with major objections because such consumption would require lactate to be a mitochondrial substrate, which requires the existence of lactate dehydrogenase in mitochondria (mLDH), an enzyme his detractors strongly insisted does not exist. Schurr and colleagues demonstrated *in vitro* that brain tissue is capable of maintaining normal neuronal function when lactate is the sole oxidizable energy substrate [16]. Skeptics of this finding argued that the phenomenon does not occur *in vivo* and even if it does, lactate cannot replace glucose as the obligatory energy substrate in brain [18]. And thus began a long-lasting debate on the validity of these findings and the potential importance of lactate as an oxidative substrate of energy metabolism in brain and elsewhere. **Figure 1** is an illustration

**33**

**Figure 1.**

*these conditions.*

*Cerebral Energy Metabolism: Measuring and Understanding Its Rate*

of the glycolytic pathway as it has been conceived and taught everywhere since 1940. This concept has been based on an initial assumption, made not by those who proposed the original reaction sequence of the glycolytic pathway, but by Krebs and Johnston, who proposed the reaction sequence of the mitochondrial TCA cycle [19]. The latter postulated that pyruvate is the substrate of that cycle and assumed that glycolysis could be its origin. Such an assumption, when made by a leading scientist of the status Krebs had achieved, was persuasive enough to compel Gustav Embden, Otto Meyerhof, and Jakub Karol Parnas to decide that pyruvate is the end-product of aerobic glycolysis and the substrate of the TCA cycle. Hence, for almost eight decades the glycolytic pathway has been described as one that has two possible outcomes, an aerobic one, which ends with pyruvate and an anaerobic one that ends with lactate (**Figure 1**). Every biochemistry textbook published from the 1940s on, every biochemistry, physiology or neuroscience course being taught at any level and every online search, all display this very description of the glycolytic pathway. That,

*The classical description of the 10 enzymatic steps of aerobic glycolysis, the pathway that converts glucose to pyruvate with a net production of two molecules of ATP and two molecules of pyruvate, its end-product and the substrate of the mitochondrial tricarboxylic acid (TCA) cycle. Under anaerobic conditions, an eleventh enzymatic step occurs where pyruvate in converted to lactate, which becomes the glycolytic end-product under* 

despite ample research data that clearly refute this dogmatic paradigm.

As indicated above, questions as to the correctness of the old description of glycolysis began to appear in the late 1980s. The lactic acidosis hypothesis of delayed neuronal damage [20] had a strong following at the time. This hypothesis postulated that lactate accumulation in the ischemic brain is the cause of delayed neuronal damage, damage observed after a recovery from the original ischemic insult had occurred. The popularity of this hypothesis was so strong that any insinuation that lactate could be anything but a menacing factor in cerebral ischemia aroused great skepticism. Consequently, when Fox et al. [21] published

**1.2 The last four decades (1980–2018)**

*DOI: http://dx.doi.org/10.5772/intechopen.84376*

*Cerebral Energy Metabolism: Measuring and Understanding Its Rate DOI: http://dx.doi.org/10.5772/intechopen.84376*

**Figure 1.**

*Cellular Metabolism and Related Disorders*

*of the brain should be perceptibly greater than that of the blood*" [1]. It was Tashiro [2] who was the first to demonstrate that nerve produces CO2 and ammonia during its metabolism. By 1924, Warburg et al. [3] demonstrated that brain tissue is able to convert large amount of glucose to lactic acid, and in 1927 it was shown that nerve produces a measurable amount of heat, an amount that increases upon electric stimulation [4]. Moreover, this increase in heat production was shown to correlate with the amount of oxygen consumed. According to Holmes [5] the above finding was the necessary proof that nerve impulse is a chemical process. All the major discoveries that have led to the elucidation of the biochemical pathways of energy metabolism were made using the tissue of choice in the field i.e., muscle. Both the tricarboxylic acid cycle and the glycolytic pathways were introduced in 1937 and 1940, respectively (the reader is directed to the many detailed reviews on the topic that are available). While there is a general agreement among biochemists, physiologists and neuroscientists as to the accuracy of the mitochondrial tricarboxylic acid (TCA) cycle and the oxidative phosphorylation pathway, disagreements exist on the accuracy of the glycolytic pathway. Hence, glycolysis is the main focus of this chapter, since the original drawing of the pathway stands in conflict with various research findings of the past three decades (for more detailed reviews of this topic the reader is directed to [6, 7]) . Although some research on brain energy metabolism was performed during 1920s and 1930s, it was limited to very few laboratories. Among them, that of Eric G. Holmes and Barbara E. Holmes pioneered important research in the field. The duo, who later joined by C.A. Ashford, published a series of papers [8–14] to demonstrate brain tissue production of lactate from glucose (similar to muscle metabolism), the involvement of phosphates and glycogen in this metabolism and the ability of brain tissue to oxidize lactate. Unfortunately, the latter never received neither the praise nor the scrutiny it deserved. This point is expanded upon in the next section, although it must be emphasized here that if the importance of that discovery would have been recognized at the time, our understanding of brain energy metabolism would be significantly accelerated and advanced. Nevertheless, Holmes and Ashford interpreted their finding of lactate oxidation simply as a process by which the brain rids itself of a waste product, since lactate was believed until the mid 1980s and even beyond, to be just that, a useless end-product of carbohydrate metabolism. Consequently, the research by Holmes and Ashford quickly became obscure. It would have stayed this way if not for a literature search I carried out in 2005 working on an upcoming paper that dealt with the possible role of lactate as an oxidative brain energy substrate [15]. Spending several weeks in the basement of my university medical school library (at the time most of the old literature has not yet been digitized) was an experience akin to treasure hunt, an experience I still cherish today. Discovering Holmes and Ashford's papers in 75-year old, dust-covered, heavy volumes of the Biochemical Journal was almost as exciting as conducting our own research [16]. Three years prior to the publication of the latter on brain energy metabolism, Brooks published his controversial work on muscle energy metabolism [17]. His proposal that skeletal muscle both produces and consumes lactate met with major objections because such consumption would require lactate to be a mitochondrial substrate, which requires the existence of lactate dehydrogenase in mitochondria (mLDH), an enzyme his detractors strongly insisted does not exist. Schurr and colleagues demonstrated *in vitro* that brain tissue is capable of maintaining normal neuronal function when lactate is the sole oxidizable energy substrate [16]. Skeptics of this finding argued that the phenomenon does not occur *in vivo* and even if it does, lactate cannot replace glucose as the obligatory energy substrate in brain [18]. And thus began a long-lasting debate on the validity of these findings and the potential importance of lactate as an oxidative substrate of energy metabolism in brain and elsewhere. **Figure 1** is an illustration

**32**

*The classical description of the 10 enzymatic steps of aerobic glycolysis, the pathway that converts glucose to pyruvate with a net production of two molecules of ATP and two molecules of pyruvate, its end-product and the substrate of the mitochondrial tricarboxylic acid (TCA) cycle. Under anaerobic conditions, an eleventh enzymatic step occurs where pyruvate in converted to lactate, which becomes the glycolytic end-product under these conditions.*

of the glycolytic pathway as it has been conceived and taught everywhere since 1940. This concept has been based on an initial assumption, made not by those who proposed the original reaction sequence of the glycolytic pathway, but by Krebs and Johnston, who proposed the reaction sequence of the mitochondrial TCA cycle [19]. The latter postulated that pyruvate is the substrate of that cycle and assumed that glycolysis could be its origin. Such an assumption, when made by a leading scientist of the status Krebs had achieved, was persuasive enough to compel Gustav Embden, Otto Meyerhof, and Jakub Karol Parnas to decide that pyruvate is the end-product of aerobic glycolysis and the substrate of the TCA cycle. Hence, for almost eight decades the glycolytic pathway has been described as one that has two possible outcomes, an aerobic one, which ends with pyruvate and an anaerobic one that ends with lactate (**Figure 1**). Every biochemistry textbook published from the 1940s on, every biochemistry, physiology or neuroscience course being taught at any level and every online search, all display this very description of the glycolytic pathway. That, despite ample research data that clearly refute this dogmatic paradigm.

#### **1.2 The last four decades (1980–2018)**

As indicated above, questions as to the correctness of the old description of glycolysis began to appear in the late 1980s. The lactic acidosis hypothesis of delayed neuronal damage [20] had a strong following at the time. This hypothesis postulated that lactate accumulation in the ischemic brain is the cause of delayed neuronal damage, damage observed after a recovery from the original ischemic insult had occurred. The popularity of this hypothesis was so strong that any insinuation that lactate could be anything but a menacing factor in cerebral ischemia aroused great skepticism. Consequently, when Fox et al. [21] published

a study, which indicated that neural activation does not require an increase in energy supply and is supported by a mere non-oxidative glucose utilization ("anaerobic" glycolysis), they met with both doubt and a degree of cynicism. Almost simultaneously, Schurr et al. [16] published their findings demonstrating that neuronal function *in vitro* can be supported by lactate as the sole oxidizable energy source. These findings were met with even greater doubt and cynicism, some of which continues to this day. The past 30 years have seen the field of brain energy metabolism grow by leaps and bounds as new technologies and techniques enable scientists to explore, measure and interpret their findings more accurately. Nevertheless, such interpretations depend on the accuracy of our knowledge and understanding of the basic pathways and processes of energy metabolism. The ongoing debate about the correct paradigm of glycolysis, as highlighted by Schurr and Gozal [22] and many of the papers within this Research Topic volume, clearly indicates that such accuracy and understanding are still to be achieved. To illustrate this point consider on one hand, the conclusion of Fox et al. [21] regarding the very low increase in energy demand upon neural stimulation, demand that can be easily answered by non-oxidative glycolysis i.e., glucose consumption unaccompanied by oxygen consumption, while on the other hand, the conclusion by Hyder et al. [23] that activated neural tissue exhibits an increase in energy production, which is fully oxidative i.e., the ratio of oxygen to glucose for this increase is 6:1. Could these completely opposing conclusions be explained by differences in the methodologies used in the two studies (the former made use of 18F-2 fluoro-2-deoxy-D-glucose to measure glucose uptake and 15O2 to measure oxygen consumption, while the latter made use of blood oxygen level dependent (BOLD) functional magnetic resonance imaging (fMRI))? Or maybe the measurements by both methods are correct, but their interpretation has relied on assumptions that emanated from older, dogmatic concept?

To answer this question, one must consider the mounting evidence supporting a paradigm shift in our comprehension of the glycolytic pathway [24]. The shift entails redrawing the glycolytic pathway as one consisting of 11 steps, beginning with glucose as its substrate and ending with lactate as its end-product, independent of the presence or absence of oxygen. From its inception, distinguishing between aerobic and anaerobic glycolysis was based not on specific evidence that the two pathways exist and produce two different products, pyruvate and lactate, respectively. That separation was an attempt by the pathway's elucidators to somehow fit it into, what they concluded, is an outcome that produces pyruvate as its main end-product since they accepted Kreb's suggestion that this monocarboxylate is the substrate of the TCA cycle. It must have been relatively easy to accept that suggestion considering lactate's negative reputation [6]. Hence, the glycolytic pathway should be considered one, uninterrupted chain of biochemical reactions that begins with glucose and ends with lactate (**Figure 1**). Accordingly, its last reaction (number 11), the reduction of pyruvate to lactate by the cytosolic lactate dehydrogenase (cLDH), plays a crucial role in keeping this pathway's cyclical nature operational as it provides a continuous supply of NAD<sup>+</sup> . If pyruvate was the glycolytic end-product, NAD<sup>+</sup> would have to be imported from other sources and locations, a proposition that has offered a somewhat shaky resolution (see [25] and references within). This, of course, is not the only factor that justifies a paradigm shift. There are numerous studies published over the past two decades demonstrating the presence of lactate dehydrogenase in mitochondria (mLDH), an enzyme that converts lactate to pyruvate [26–32]. Brooks et al. [33] also demonstrated the presence of monocarboxylate transporter 1 (MCT1) in mitochondria, the transporter that is responsible for the transport of lactate along its gradient from the cytosol to the mitochondrion. Havel et al. showed that in blood and in other tissues the

**35**

**Figure 2.**

*the oxygenation conditions (C).*

*Cerebral Energy Metabolism: Measuring and Understanding Its Rate*

ratio lactate/pyruvate is >10, a value that is not consistent with the assumption that pyruvate is the glycolytic end-product [34]. Moreover, the proposal that aerobic glycolysis ends with pyruvate does not meet the known standard free-energy (∆G0') change of the reaction pyruvate → lactate, which is −6.0 kcal/mol, a value indicating that this reaction should proceed independently of the presence or absence of oxygen. In other words, glycolysis, whether aerobic or anaerobic, should always end up with lactate. **Figure 2A** demonstrates the free energy change profile of aerobic glycolysis that ends with the reaction phosphoenolpyruvate → pyruvate, although the potential free-energy change of the conversion pyruvate → lactate (**Figure 2B**, anaerobic glycolysis) determines that glycolysis should end with lactate regardless of the oxygenation condition (**Figure 2C**). Last but not least, the reaction equilibrium of cLDH is tilted heavily in the direction of lactate production, which makes it unlikely for lactate to be converted back to pyruvate by that cytosolic enzyme. In contrast, the reaction equilibrium of mLDH tilts in the direction of lactate oxida-

The above points support the proposed paradigm shift in the glycolytic pathway [24], where lactate, not pyruvate, is its end-product and the oxidative mitochondrial substrate for the TCA cycle. Accordingly, is measuring the cerebral

brain's ability to handle its energy demands under those conditions?

an accurate picture of brain energy metabolism during rest or activation, in health or disease? If lactate is an oxidative energy substrate, should not CMRlactate also be measured in order to have a more complete account of cerebral energy metabolism? How would the measurement of CMRlactate contribute to our understanding of the

*A schematic illustration of the potential free-energy change profile of aerobic (A) and anaerobic glycolysis (B). The potential free-energy change of the reaction pyruvate → lactate dictates that it should proceed regardless of* 

) and glucose (CMRglucose) sufficient in providing

*DOI: http://dx.doi.org/10.5772/intechopen.84376*

tion to pyruvate [35, 36].

metabolic rates of oxygen (CMRO2

#### *Cerebral Energy Metabolism: Measuring and Understanding Its Rate DOI: http://dx.doi.org/10.5772/intechopen.84376*

*Cellular Metabolism and Related Disorders*

emanated from older, dogmatic concept?

tional as it provides a continuous supply of NAD<sup>+</sup>

a study, which indicated that neural activation does not require an increase in energy supply and is supported by a mere non-oxidative glucose utilization ("anaerobic" glycolysis), they met with both doubt and a degree of cynicism. Almost simultaneously, Schurr et al. [16] published their findings demonstrating that neuronal function *in vitro* can be supported by lactate as the sole oxidizable energy source. These findings were met with even greater doubt and cynicism, some of which continues to this day. The past 30 years have seen the field of brain energy metabolism grow by leaps and bounds as new technologies and techniques enable scientists to explore, measure and interpret their findings more accurately. Nevertheless, such interpretations depend on the accuracy of our knowledge and understanding of the basic pathways and processes of energy metabolism. The ongoing debate about the correct paradigm of glycolysis, as highlighted by Schurr and Gozal [22] and many of the papers within this Research Topic volume, clearly indicates that such accuracy and understanding are still to be achieved. To illustrate this point consider on one hand, the conclusion of Fox et al. [21] regarding the very low increase in energy demand upon neural stimulation, demand that can be easily answered by non-oxidative glycolysis i.e., glucose consumption unaccompanied by oxygen consumption, while on the other hand, the conclusion by Hyder et al. [23] that activated neural tissue exhibits an increase in energy production, which is fully oxidative i.e., the ratio of oxygen to glucose for this increase is 6:1. Could these completely opposing conclusions be explained by differences in the methodologies used in the two studies (the former made use of 18F-2 fluoro-2-deoxy-D-glucose to measure glucose uptake and 15O2 to measure oxygen consumption, while the latter made use of blood oxygen level dependent (BOLD) functional magnetic resonance imaging (fMRI))? Or maybe the measurements by both methods are correct, but their interpretation has relied on assumptions that

To answer this question, one must consider the mounting evidence supporting a paradigm shift in our comprehension of the glycolytic pathway [24]. The shift entails redrawing the glycolytic pathway as one consisting of 11 steps, beginning with glucose as its substrate and ending with lactate as its end-product, independent of the presence or absence of oxygen. From its inception, distinguishing between aerobic and anaerobic glycolysis was based not on specific evidence that the two pathways exist and produce two different products, pyruvate and lactate, respectively. That separation was an attempt by the pathway's elucidators to somehow fit it into, what they concluded, is an outcome that produces pyruvate as its main end-product since they accepted Kreb's suggestion that this monocarboxylate is the substrate of the TCA cycle. It must have been relatively easy to accept that suggestion considering lactate's negative reputation [6]. Hence, the glycolytic pathway should be considered one, uninterrupted chain of biochemical reactions that begins with glucose and ends with lactate (**Figure 1**). Accordingly, its last reaction (number 11), the reduction of pyruvate to lactate by the cytosolic lactate dehydrogenase (cLDH), plays a crucial role in keeping this pathway's cyclical nature opera-

. If pyruvate was the glycolytic

would have to be imported from other sources and locations,

a proposition that has offered a somewhat shaky resolution (see [25] and references within). This, of course, is not the only factor that justifies a paradigm shift. There are numerous studies published over the past two decades demonstrating the presence of lactate dehydrogenase in mitochondria (mLDH), an enzyme that converts lactate to pyruvate [26–32]. Brooks et al. [33] also demonstrated the presence of monocarboxylate transporter 1 (MCT1) in mitochondria, the transporter that is responsible for the transport of lactate along its gradient from the cytosol to the mitochondrion. Havel et al. showed that in blood and in other tissues the

**34**

end-product, NAD<sup>+</sup>

ratio lactate/pyruvate is >10, a value that is not consistent with the assumption that pyruvate is the glycolytic end-product [34]. Moreover, the proposal that aerobic glycolysis ends with pyruvate does not meet the known standard free-energy (∆G0') change of the reaction pyruvate → lactate, which is −6.0 kcal/mol, a value indicating that this reaction should proceed independently of the presence or absence of oxygen. In other words, glycolysis, whether aerobic or anaerobic, should always end up with lactate. **Figure 2A** demonstrates the free energy change profile of aerobic glycolysis that ends with the reaction phosphoenolpyruvate → pyruvate, although the potential free-energy change of the conversion pyruvate → lactate (**Figure 2B**, anaerobic glycolysis) determines that glycolysis should end with lactate regardless of the oxygenation condition (**Figure 2C**). Last but not least, the reaction equilibrium of cLDH is tilted heavily in the direction of lactate production, which makes it unlikely for lactate to be converted back to pyruvate by that cytosolic enzyme. In contrast, the reaction equilibrium of mLDH tilts in the direction of lactate oxidation to pyruvate [35, 36].

The above points support the proposed paradigm shift in the glycolytic pathway [24], where lactate, not pyruvate, is its end-product and the oxidative mitochondrial substrate for the TCA cycle. Accordingly, is measuring the cerebral metabolic rates of oxygen (CMRO2 ) and glucose (CMRglucose) sufficient in providing an accurate picture of brain energy metabolism during rest or activation, in health or disease? If lactate is an oxidative energy substrate, should not CMRlactate also be measured in order to have a more complete account of cerebral energy metabolism? How would the measurement of CMRlactate contribute to our understanding of the brain's ability to handle its energy demands under those conditions?

#### **Figure 2.**

*A schematic illustration of the potential free-energy change profile of aerobic (A) and anaerobic glycolysis (B). The potential free-energy change of the reaction pyruvate → lactate dictates that it should proceed regardless of the oxygenation conditions (C).*

#### **2. Measurement of cerebral energy metabolic rates**

At the basis of each technology designed to measure the rate of brain energy metabolism is the idea that measuring the consumption rate of the main two substrates of glycolysis and mitochondrial respiration, glucose and oxygen (O2), should provide a complete picture of the brain's energy use. Theoretically, under normal conditions, each glucose molecule that enters the glycolytic pathway requires six molecules of oxygen to be fully oxidized via the mitochondrial TCA cycle and the electron transport chain. Thus, simultaneous measurements of glucose and oxygen consumption during rest or activation supposedly produces accurate estimate of the energy needs for the brain region under observation. However, the ratio CMRO2 /CMRglucose values calculated are often significantly smaller than the expected 6/1. Such discrepancies have attributed to other glucose-consuming reactions not accompanies with oxygen consumption. Consequently, it has been a common understanding that a value of CMRO2 /CMRglucose < 6 indicates that a partial non-oxidative glucose consumption. The smaller the value of CMRO2 /CMRglucose, the greater is the non-oxidative consumption of glucose. This understanding makes sense when one assumes that a fully coupled glycolytic-mitochondrial respiratory apparatus should produce a CMRO2 /CMRglucose value of 6 and an uncoupled apparatus (non-oxidative) should produce a CMRO2 /CMRglucose value of ~0.

As indicated above, myriad techniques and technologies have been developed during the past six decades to measure both CMRO2 and CMRglucose. To measure cerebral energy metabolism *in vivo* one can analyze chemical changes in the blood entering and exiting the brain and/or in the cerebrospinal fluid. Of course, brain tissue samples can also be taken for analysis before and after physiological activity, although this approach would lend itself only to experimental animals. The introduction of radioisotopes to the analytical techniques of brain metabolic activity has greatly improved their speed and accuracy. Radioisotopes allow not only the tracing of end-products of cerebral metabolism, but also the detection of intermediates of that metabolism. Nevertheless, these techniques have their own drawbacks, including the need to sacrifice the animal under study only to receive a single measurement which provides mainly a qualitative value. A quantitative measurement is frequently confounded by compartmentation and its misinterpretation thereof. One of the most reliable techniques to measure oxygen consumption is the polarographic technique, which allows the determination of oxygen concentration via the measurement of the partial oxygen pressure (PO2) locally. Continuous measurements over a period of time when brain activity (EEG) is monitored, demonstrated a correlation between increased activity and decreased tissue oxygen level. The development of oxygen microelectrodes has afforded a more accurate localization of such measurements.

In principle, CMR can be expressed as: CMR = CBF (A − V).

where (A − V) is the difference in concentration in arterial and cerebral venous blood, CBF is the rate of cerebral blood flow in volume of blood per unit time, a CMR (cerebral metabolic rate) is the steady state of utilization or production of a substance by the brain [18]. This equation is the foundation on which quantitative CMR studies *in vivo* have been conducted. Since the normal brain consumed approximately 20% of the total body oxygen consumption to maintain its functionality and structure, it is clear that any interruption in this high demand for oxidative energy metabolism could have far reaching survivability consequences. Clearly, a nonoxidative energy metabolism (glycolysis) is incapable of answering the high energy demands of the brain. That a stimulated brain has still higher energy demands than the resting one would be an inevitable conclusion. Hence, when studying the energy demands of a specific activated brain region, such activation is expected to produce

**37**

*Cerebral Energy Metabolism: Measuring and Understanding Its Rate*

their study under the title "Nonoxidative glucose consumption during focal physiologic neural activity" [21] they stirred a small tempest among scientists in the community that studies cerebral blood flow and metabolism. These investigators employed 18F-labeled 2-fluoro-2-deoxy-D-glucose to measure CMRglucose, a method originally developed over a decade earlier [37], and 15O-labeled molecular O2 to mea-

glucose tissue uptake in excess of that consumed by oxidative metabolism. They concluded their findings to indicate that stimulated brain activity requires significantly less energy than previously thought. Also, since they measured a corresponding increase in CBF along with the increase in glucose consumption, the investigators argued that this increase is for purposes other than oxidative metabolism. These conclusions stemmed from the prevailing postulate that over 90% of resting brain's glucose consumption is oxidative and less than 5% of that consumption ends in glycolytic lactate production. Since the oxidative consumption of one molecule of glucose produces approximately 36 molecules of ATP, while the glycolytic consumption of one molecule of glucose produces only 2 molecules of ATP, one can easily appreciate how oxidative consumption of glucose is responsible for 90% of the resting brain ATP production. Hence, the finding by [21] that brain stimulation increased glucose consumption without a corresponding increase in oxygen consumption unsettled the established understanding according to which increased brain activity must be accompanied by a corresponding increase in energy supply. This seminal paper was originated from the laboratory of Marcus Raichle, a laboratory that has become a leading center for functional brain imaging [38]. Imaging technologies such as X-ray computed tomography (CT), positron emission tomography (PET), near-infrared spectroscopy (NIRS) and magnetic resonance imaging (MRI) are the main techniques available for the measurement of brain energy metabolism during rest and activity. The most popular technology for this purpose today is the blood oxygen level dependent (BOLD) functional magnetic resonance imaging (fMRI), which was developed by [39]. BOLD fMRI measures changes in blood oxygenation in relation to brain activity, although that relationship is somewhat ambiguous, since it is not accompanied by a direct neural activity measurement such as that allowed by electrophysiology. When the latter is combined with direct oxygen concentration measurements, using an oxygen microelectrode (polarography), a higher resolution than BOLD fMRI can be achieved [40]. Besides tissue oxygen measurements using microelectrodes, tissue glucose and lactate concentrations can also be assessed using specific microelectrodes (sensors). Of course, this approach does not lend itself for regular use in humans, however, for the purpose of *in vivo* studies in experimental

animals, the approach proved itself to be very useful and an eye opener.

In this respect, Hu and Wilson [41] studied the coupling of a temporary local energy pool to neuronal activity in the rat brain (**Figure 3**). They were the first to combine the use of three separate rapid response sensors (microelectrodes) to measure tissue oxygen, glucose and lactate concentrations. The investigators placed them in the dentate gyrus of the rat hippocampus, observing how they fluctuate in response to 10 consecutive electrical stimulations of the perforant pathway (each stimulus lasted 5 s and applied every 2 min). Their results were analyzed by Schurr and Gozal [36] (**Figure 3**). A literature search shows that Hu and Wilson's interpretation of their findings has its supporters [15, 42–47] and detractors [48–50]. The former group argued that these findings are strengthening the concept that lactate is the energy substrate that is utilized oxidatively upon neuronal activation. The latter group disagreed with this conclusion. Schurr [24] further analyzed the results of Hu and Wilson [41] beyond an earlier analysis [36]. The more recent analysis was prompted for two reasons. First, two decades have passed since the publication

. They demonstrated that transient increases in neural activity elevated

and CMRglucose. Consequently, when Fox et al. published

*DOI: http://dx.doi.org/10.5772/intechopen.84376*

an increase in both CMRO2

sure CMRO2

#### *Cerebral Energy Metabolism: Measuring and Understanding Its Rate DOI: http://dx.doi.org/10.5772/intechopen.84376*

*Cellular Metabolism and Related Disorders*

the ratio CMRO2

**2. Measurement of cerebral energy metabolic rates**

common understanding that a value of CMRO2

tus (non-oxidative) should produce a CMRO2

during the past six decades to measure both CMRO2

apparatus should produce a CMRO2

of such measurements.

At the basis of each technology designed to measure the rate of brain energy metabolism is the idea that measuring the consumption rate of the main two substrates of glycolysis and mitochondrial respiration, glucose and oxygen (O2), should provide a complete picture of the brain's energy use. Theoretically, under normal conditions, each glucose molecule that enters the glycolytic pathway requires six molecules of oxygen to be fully oxidized via the mitochondrial TCA cycle and the electron transport chain. Thus, simultaneous measurements of glucose and oxygen consumption during rest or activation supposedly produces accurate estimate of the energy needs for the brain region under observation. However,

the expected 6/1. Such discrepancies have attributed to other glucose-consuming reactions not accompanies with oxygen consumption. Consequently, it has been a

the greater is the non-oxidative consumption of glucose. This understanding makes sense when one assumes that a fully coupled glycolytic-mitochondrial respiratory

As indicated above, myriad techniques and technologies have been developed

cerebral energy metabolism *in vivo* one can analyze chemical changes in the blood entering and exiting the brain and/or in the cerebrospinal fluid. Of course, brain tissue samples can also be taken for analysis before and after physiological activity, although this approach would lend itself only to experimental animals. The introduction of radioisotopes to the analytical techniques of brain metabolic activity has greatly improved their speed and accuracy. Radioisotopes allow not only the tracing of end-products of cerebral metabolism, but also the detection of intermediates of that metabolism. Nevertheless, these techniques have their own drawbacks, including the need to sacrifice the animal under study only to receive a single measurement which provides mainly a qualitative value. A quantitative measurement is frequently confounded by compartmentation and its misinterpretation thereof. One of the most reliable techniques to measure oxygen consumption is the polarographic technique, which allows the determination of oxygen concentration via the measurement of the partial oxygen pressure (PO2) locally. Continuous measurements over a period of time when brain activity (EEG) is monitored, demonstrated a correlation between increased activity and decreased tissue oxygen level. The development of oxygen microelectrodes has afforded a more accurate localization

non-oxidative glucose consumption. The smaller the value of CMRO2

In principle, CMR can be expressed as: CMR = CBF (A − V).

where (A − V) is the difference in concentration in arterial and cerebral venous blood, CBF is the rate of cerebral blood flow in volume of blood per unit time, a CMR (cerebral metabolic rate) is the steady state of utilization or production of a substance by the brain [18]. This equation is the foundation on which quantitative CMR studies *in vivo* have been conducted. Since the normal brain consumed approximately 20% of the total body oxygen consumption to maintain its functionality and structure, it is clear that any interruption in this high demand for oxidative energy metabolism could have far reaching survivability consequences. Clearly, a nonoxidative energy metabolism (glycolysis) is incapable of answering the high energy demands of the brain. That a stimulated brain has still higher energy demands than the resting one would be an inevitable conclusion. Hence, when studying the energy demands of a specific activated brain region, such activation is expected to produce

/CMRglucose values calculated are often significantly smaller than

/CMRglucose < 6 indicates that a partial

and CMRglucose. To measure

/CMRglucose value of 6 and an uncoupled appara-

/CMRglucose value of ~0.

/CMRglucose,

**36**

an increase in both CMRO2 and CMRglucose. Consequently, when Fox et al. published their study under the title "Nonoxidative glucose consumption during focal physiologic neural activity" [21] they stirred a small tempest among scientists in the community that studies cerebral blood flow and metabolism. These investigators employed 18F-labeled 2-fluoro-2-deoxy-D-glucose to measure CMRglucose, a method originally developed over a decade earlier [37], and 15O-labeled molecular O2 to measure CMRO2 . They demonstrated that transient increases in neural activity elevated glucose tissue uptake in excess of that consumed by oxidative metabolism. They concluded their findings to indicate that stimulated brain activity requires significantly less energy than previously thought. Also, since they measured a corresponding increase in CBF along with the increase in glucose consumption, the investigators argued that this increase is for purposes other than oxidative metabolism. These conclusions stemmed from the prevailing postulate that over 90% of resting brain's glucose consumption is oxidative and less than 5% of that consumption ends in glycolytic lactate production. Since the oxidative consumption of one molecule of glucose produces approximately 36 molecules of ATP, while the glycolytic consumption of one molecule of glucose produces only 2 molecules of ATP, one can easily appreciate how oxidative consumption of glucose is responsible for 90% of the resting brain ATP production. Hence, the finding by [21] that brain stimulation increased glucose consumption without a corresponding increase in oxygen consumption unsettled the established understanding according to which increased brain activity must be accompanied by a corresponding increase in energy supply. This seminal paper was originated from the laboratory of Marcus Raichle, a laboratory that has become a leading center for functional brain imaging [38]. Imaging technologies such as X-ray computed tomography (CT), positron emission tomography (PET), near-infrared spectroscopy (NIRS) and magnetic resonance imaging (MRI) are the main techniques available for the measurement of brain energy metabolism during rest and activity. The most popular technology for this purpose today is the blood oxygen level dependent (BOLD) functional magnetic resonance imaging (fMRI), which was developed by [39]. BOLD fMRI measures changes in blood oxygenation in relation to brain activity, although that relationship is somewhat ambiguous, since it is not accompanied by a direct neural activity measurement such as that allowed by electrophysiology. When the latter is combined with direct oxygen concentration measurements, using an oxygen microelectrode (polarography), a higher resolution than BOLD fMRI can be achieved [40]. Besides tissue oxygen measurements using microelectrodes, tissue glucose and lactate concentrations can also be assessed using specific microelectrodes (sensors). Of course, this approach does not lend itself for regular use in humans, however, for the purpose of *in vivo* studies in experimental animals, the approach proved itself to be very useful and an eye opener.

In this respect, Hu and Wilson [41] studied the coupling of a temporary local energy pool to neuronal activity in the rat brain (**Figure 3**). They were the first to combine the use of three separate rapid response sensors (microelectrodes) to measure tissue oxygen, glucose and lactate concentrations. The investigators placed them in the dentate gyrus of the rat hippocampus, observing how they fluctuate in response to 10 consecutive electrical stimulations of the perforant pathway (each stimulus lasted 5 s and applied every 2 min). Their results were analyzed by Schurr and Gozal [36] (**Figure 3**). A literature search shows that Hu and Wilson's interpretation of their findings has its supporters [15, 42–47] and detractors [48–50]. The former group argued that these findings are strengthening the concept that lactate is the energy substrate that is utilized oxidatively upon neuronal activation. The latter group disagreed with this conclusion. Schurr [24] further analyzed the results of Hu and Wilson [41] beyond an earlier analysis [36]. The more recent analysis was prompted for two reasons. First, two decades have passed since the publication

#### **Figure 3.**

*Profiles of time course and dynamic relationships of local extracellular lactate, glucose, and PO2 levels in the rat hippocampal dentate gyrus during a series of 5 s electrical stimulations (arrows) of the perforant pathway at 2 min rest intervals (reproduced with permission from Hu and Wilson, copyright 1997, Blackwell, Oxford). The changes in the mean concentration of glucose were always in opposite direction to the changes in mean lactate concentration. The vertical lines were drawn to indicate the simultaneous dip in all three analytes in response to each of the electrical stimulations. For additional details see [41] from where the figure and the legend have been reproduced with permission and [36].*

of the paper by Hu and Wilson [41], a period in which numerous studies added much support to the idea that lactate is a mitochondrial oxidative energy substrate. Second, many other studies on cerebral energy metabolism continue to conclude that neural activity is supported by "anaerobic" glycolysis and not by oxidative utilization of glucose, while ignoring the possibility that such activity may be supported by oxidative utilization of lactate.

#### **3. Lactate cerebral metabolic rate and the importance of its measurement**

When the results of the study by Hu and Wilson [41] were analyzed before [36], the analysis showed that upon a series of 10 stimulation of the rat hippocampal perforant pathway a steady glucose consumption was accompanied by a gradual increase in lactate consumption. Considering the conclusion of Fox et al. [21] that aerobic glycolytic ATP production is sufficient to answer the energy needs of activated neural tissue, one could assume that it should be sufficient to provide the energy needs of the stimulated hippocampal dentate gyrus. In addition, this analysis points out that if the conclusion of Fox et al. [21] is correct, the energy needs of the activated dentate gyrus declined with each stimulation or stayed the same at a very low level of ATP production (0.8–0.3 mM). However, if lactate oxidative consumption is postulated to be responsible for the ATP production that sustains the energy needs of the stimulated tissue, the increased lactate consumption with each consecutive stimulation signals a concomitant increased ATP production. The calculation shows that the response to the first stimulation produced 3 mM ATP, while the response to the last stimulation produced almost 11 mM (**Figure 4**).

The more recent analysis [24] also indicates that the increased levels of tissue lactate following each stimulation [41] could not be produced from the glycolytically metabolized glucose (**Figure 5**). Hence, this additional lactate had to be originated from other sources i.e., the surrounding tissue or glycogen stores [50]. As was shown by Hu and Wilson [41](**Figure 3**), a larger amount of lactate was consumed during each consecutive stimulation, while a smaller amount of glucose was

**39**

**Figure 4.**

*Cerebral Energy Metabolism: Measuring and Understanding Its Rate*

consumed. Moreover, following each stimulation, except the first one, the lactate level measured exceeded the level expected from the amount of glucose consumed glycolytically i.e., two moles of lactate from one mole of glucose. Following the second stimulation, the tissue ratio of lactate to glucose was 3.95 and by the 10th stimulation this ratio increased to 8.33 (**Figure 5**). Meanwhile, oxygen tissue levels dipped and rose as expected during and after each stimulation, respectively, signaling that the electrical stimulation evoked an oxidative consumption of substrate. Initially, glucose and lactate were oxidatively consumed at equal amounts however,

*Time course of changes in the amplitude of the dip in tissue glucose and lactate levels in the rat hippocampal dentate gyrus after each of the 10 electrical stimulations applied to the perforant pathway at intervals of 2 min (bottom panel). The amplitude of each dip (in mM) was calculated from the Hu and Wilson [41] as reproduced in Figure 3. The upper panel represents the estimated ATP amount produced based on the size of the dip (in mM) in tissue glucose and lactate levels as shown in the bottom panel. The estimated ATP levels were calculated as follows: the glucose measured dip (in mM) was multiplied by 2, the net production of 2 moles ATP from each mole of glucose metabolized glycolytically; the lactate measure dip (in mM) was multiplied by 34, the net formation of 34 moles of ATP for every 2 moles of lactate (form glycolytically from 1 mole of glucose) metabolized via the mitochondrial TCA cycle and the oxidative phosphorylation chain.*

*DOI: http://dx.doi.org/10.5772/intechopen.84376*

#### **Figure 4.**

*Cellular Metabolism and Related Disorders*

by oxidative utilization of lactate.

*legend have been reproduced with permission and [36].*

**measurement**

**Figure 3.**

of the paper by Hu and Wilson [41], a period in which numerous studies added much support to the idea that lactate is a mitochondrial oxidative energy substrate. Second, many other studies on cerebral energy metabolism continue to conclude that neural activity is supported by "anaerobic" glycolysis and not by oxidative utilization of glucose, while ignoring the possibility that such activity may be supported

*Profiles of time course and dynamic relationships of local extracellular lactate, glucose, and PO2 levels in the rat hippocampal dentate gyrus during a series of 5 s electrical stimulations (arrows) of the perforant pathway at 2 min rest intervals (reproduced with permission from Hu and Wilson, copyright 1997, Blackwell, Oxford). The changes in the mean concentration of glucose were always in opposite direction to the changes in mean lactate concentration. The vertical lines were drawn to indicate the simultaneous dip in all three analytes in response to each of the electrical stimulations. For additional details see [41] from where the figure and the* 

When the results of the study by Hu and Wilson [41] were analyzed before [36],

the analysis showed that upon a series of 10 stimulation of the rat hippocampal perforant pathway a steady glucose consumption was accompanied by a gradual increase in lactate consumption. Considering the conclusion of Fox et al. [21] that aerobic glycolytic ATP production is sufficient to answer the energy needs of activated neural tissue, one could assume that it should be sufficient to provide the energy needs of the stimulated hippocampal dentate gyrus. In addition, this analysis points out that if the conclusion of Fox et al. [21] is correct, the energy needs of the activated dentate gyrus declined with each stimulation or stayed the same at a very low level of ATP production (0.8–0.3 mM). However, if lactate oxidative consumption is postulated to be responsible for the ATP production that sustains the energy needs of the stimulated tissue, the increased lactate consumption with each consecutive stimulation signals a concomitant increased ATP production. The calculation shows that the response to the first stimulation produced 3 mM ATP, while the response to the last stimulation produced almost 11 mM (**Figure 4**). The more recent analysis [24] also indicates that the increased levels of tissue lactate following each stimulation [41] could not be produced from the glycolytically metabolized glucose (**Figure 5**). Hence, this additional lactate had to be originated from other sources i.e., the surrounding tissue or glycogen stores [50]. As was shown by Hu and Wilson [41](**Figure 3**), a larger amount of lactate was consumed during each consecutive stimulation, while a smaller amount of glucose was

**3. Lactate cerebral metabolic rate and the importance of its** 

**38**

*Time course of changes in the amplitude of the dip in tissue glucose and lactate levels in the rat hippocampal dentate gyrus after each of the 10 electrical stimulations applied to the perforant pathway at intervals of 2 min (bottom panel). The amplitude of each dip (in mM) was calculated from the Hu and Wilson [41] as reproduced in Figure 3. The upper panel represents the estimated ATP amount produced based on the size of the dip (in mM) in tissue glucose and lactate levels as shown in the bottom panel. The estimated ATP levels were calculated as follows: the glucose measured dip (in mM) was multiplied by 2, the net production of 2 moles ATP from each mole of glucose metabolized glycolytically; the lactate measure dip (in mM) was multiplied by 34, the net formation of 34 moles of ATP for every 2 moles of lactate (form glycolytically from 1 mole of glucose) metabolized via the mitochondrial TCA cycle and the oxidative phosphorylation chain.*

consumed. Moreover, following each stimulation, except the first one, the lactate level measured exceeded the level expected from the amount of glucose consumed glycolytically i.e., two moles of lactate from one mole of glucose. Following the second stimulation, the tissue ratio of lactate to glucose was 3.95 and by the 10th stimulation this ratio increased to 8.33 (**Figure 5**). Meanwhile, oxygen tissue levels dipped and rose as expected during and after each stimulation, respectively, signaling that the electrical stimulation evoked an oxidative consumption of substrate. Initially, glucose and lactate were oxidatively consumed at equal amounts however,

#### **Figure 5.**

*The local extracellular glucose, lactate and O2 levels in a rat hippocampal dentate gyrus during a series of 5 s electrical stimulations of the perforant pathway at 2 min rest intervals and the dynamic relationship between them. Glucose, lactate and O2 concentrations were calculated from their dips and rises as measured by Hu and Wilson [41] using rapid response sensors. The numerical values posted above the columns representing the rises in glucose and lactate post-stimulation are the calculated ratios between the two. For additional details see Figure 3 and [36].*

from the second stimulus onward more lactate than glucose was consumed (**Figure 5**). The oxygen level as measured by Hu and Wilson [41] fluctuated with a dip upon stimulation and a sharp rise upon its cessation (**Figure 3**). The fast rise can be interpreted as evidence that ample oxygen was available if and when needed. This rise also indicates that the tissue was well oxygenated during the duration of the experiment. Considering that one mole of lactate consumes three moles of oxygen for its full oxidation as compared to six moles of oxygen consumed by glucose for its full oxidation, if lactate, rather than glucose, is the main oxidative energy substrate during neural tissue activation, the expected ratio CMR <sup>O</sup> <sup>2</sup> :CMRlactate should not exceed 3:1. Therefore, it is reasonable to presume that during neural activation, when lactate oxidation is a major supplier of the ATP necessary to support said activation, the ratio CMR <sup>O</sup> <sup>2</sup> :CMRglucose should be considerably lower than 6:1. Obviously, most, if not all, studies aimed at measuring cerebral metabolic rates postulate that the ratio CMR <sup>O</sup> <sup>2</sup> :CMRglucose measured or calculated should approach 6 [23]. The conclusions of Fox et al. [21] are in complete disagreement with the measurements and calculations of Hyder et al. [23]. While the former concludes an almost complete uncoupling between glucose and oxygen consumption by activated neural tissue, the latter asserts the maintenance of full coupling between glucose and oxygen consumption during neural activation. In both studies [21, 23] the interpretation of the results is based entirely on the original, dogmatic paradigm of glycolysis according to which aerobic glycolysis ends with pyruvate, the assumed substrate of the mitochondrial TCA cycle.

The *in vivo* measurements performed by Hu and Wilson [41] of both glucose and lactate concentrations before, during and post electrical stimulation provide much support to the proposal that lactate plays a major role in oxidative energy metabolism of the activated neural tissue. The relatively small dips and rises in O2 levels in response to the electrical stimulations, as measured polarographically, should be given additional consideration. First, the spatial and temporal resolutions provided by polarographic measurement of O2 compared to BOLD fMRI measurements allow for a better characterization of the time-course of oxygen responses [40].

**41**

*Cerebral Energy Metabolism: Measuring and Understanding Its Rate*

While BOLD fMRI estimates yielded a CMRO2

Hyder et al. [23] used BOLD fMRI and the measurements used by Fox et al. [21] were even more cumbersome, involving the use of [15O]H2O, [15O]O2 and [15O]CO2.

measurements produced a ratio value of 0.4:1. These completely opposing outcomes make one wonder whether or not the measurements performed using these two

in the consumption of molecular oxygen upon neural activation. Could the direct

with the indirect ones made by Fox et al. [21] and Hyder et al. [23], be reconciled such that a better picture of cerebral metabolic rates of activated neural tissue can be visualized? It is widely agreed that over 90% of the normal brain's energy production originates from glucose oxidation [21, 51]. The normal glucose concentration in the brain is ~2 mM and its normal lactate concentration is about half of that of glucose. Thus, it is safe to postulate that the normal resting brain is supplied with ample amounts of oxygen to continuously oxidize more than 90% of the brain glucose. However, glucose supplies to the normal brain are limited (only 40% of normal blood glucose level). Consequently, the increased rate of CBF along with the increased consumption of glucose upon activation [21, 23, 52] should supply all the oxygen necessary to match the increased demand, in contrast to the limited supplies of glucose. Low resolution techniques for the measurements of oxygen concentrations are unable to detect local fluctuations accurately if at all, which could explain why Fox et al. [21] reached the conclusion regarding the very low oxygen consumption during neural activation. Nonetheless, their conclusion that the energy demands of activated neural tissue are being met through glycolytic ATP production is most likely incorrect. In other words, undetectable or slightly detectable dip in tissue oxygen level upon activation is not necessarily an indication that oxygen is not consumed. The higher resolution of oxygen measurement afforded by polarography exemplifies the fact that local oxygen levels dipped upon stimulation and overshot upon its cessation ([41]; **Figures 3** and **4**). Although local fluctuations in tissue oxygen levels were evident, its overall tissue concentration did not significantly change and may even have risen somewhat above its baseline level. In contrast, both glucose and lactate levels were changed significantly from their baseline levels [24, 36, 41] (**Figures 3**–**5**). The fluctuations between lactate and oxygen were highly synchronized, indicating that lactate is being oxidized upon tissue activation. During the 20 min following the 10th stimulation, the tissue level of both oxygen and glucose climbed above the baseline level, while the high level of lactate gradually declined ([41]; **Figure 3**). These shifts seem to indicate that upon cessation of stimulation, as the tissue is recovering from activation and high energy demands, lactate becomes the preferred oxidative energy substrate, sparing glucose. That the cerebral tissue would prefer lactate over glucose, especially when the former is abundant, is reasonable, considering the fact that lactate oxidative mitochondria. Consequently, the increased rate of CBF along with the increased consumption of glucose upon activation [21, 23, 52] should supply all the oxygen necessary to match the increased demand, in contrast to the limited supplies of glucose. Low resolution techniques for the measurements of oxygen concentrations are unable to detect local fluctuations accurately if at all, which could explain why Fox et al. [21] reached the conclusion regarding the very low oxygen consumption during neural activation. Nonetheless, their conclusion that the energy demands of activated neural tissue are being met through glycolytic ATP production is most likely incorrect. In other words, undetectable or slightly detectable dip in tissue oxygen level upon activation is not necessarily an indication that oxygen is not consumed. The higher resolution of oxygen measurement afforded by polarography exemplifies the fact that local oxygen levels dipped upon stimulation and overshot upon its cessation (**Figures 3** and **4** and [41]). Although local fluctuations in tissue

:CMRglucose ratio value of 6:1, the 15O

they produced truly reflect the changes

, CMRglucose and CMRlactate done by Hu and Wilson [41] along

*DOI: http://dx.doi.org/10.5772/intechopen.84376*

methods and the calculated values of CMRO2

measurements of CMRO2

#### *Cerebral Energy Metabolism: Measuring and Understanding Its Rate DOI: http://dx.doi.org/10.5772/intechopen.84376*

*Cellular Metabolism and Related Disorders*

from the second stimulus onward more lactate than glucose was consumed (**Figure 5**). The oxygen level as measured by Hu and Wilson [41] fluctuated with a dip upon stimulation and a sharp rise upon its cessation (**Figure 3**). The fast rise can be interpreted as evidence that ample oxygen was available if and when needed. This rise also indicates that the tissue was well oxygenated during the duration of the experiment. Considering that one mole of lactate consumes three moles of oxygen for its full oxidation as compared to six moles of oxygen consumed by glucose for its full oxidation, if lactate, rather than glucose, is the main oxidative energy substrate dur-

*The local extracellular glucose, lactate and O2 levels in a rat hippocampal dentate gyrus during a series of 5 s electrical stimulations of the perforant pathway at 2 min rest intervals and the dynamic relationship between them. Glucose, lactate and O2 concentrations were calculated from their dips and rises as measured by Hu and Wilson [41] using rapid response sensors. The numerical values posted above the columns representing the rises in glucose and lactate post-stimulation are the calculated ratios between the two. For additional details see* 

3:1. Therefore, it is reasonable to presume that during neural activation, when lactate oxidation is a major supplier of the ATP necessary to support said activation,

if not all, studies aimed at measuring cerebral metabolic rates postulate that the

sions of Fox et al. [21] are in complete disagreement with the measurements and calculations of Hyder et al. [23]. While the former concludes an almost complete uncoupling between glucose and oxygen consumption by activated neural tissue, the latter asserts the maintenance of full coupling between glucose and oxygen consumption during neural activation. In both studies [21, 23] the interpretation of the results is based entirely on the original, dogmatic paradigm of glycolysis according to which aerobic glycolysis ends with pyruvate, the assumed substrate of the

The *in vivo* measurements performed by Hu and Wilson [41] of both glucose and lactate concentrations before, during and post electrical stimulation provide much support to the proposal that lactate plays a major role in oxidative energy metabolism of the activated neural tissue. The relatively small dips and rises in O2 levels in response to the electrical stimulations, as measured polarographically, should be given additional consideration. First, the spatial and temporal resolutions provided by polarographic measurement of O2 compared to BOLD fMRI measurements allow

for a better characterization of the time-course of oxygen responses [40].

:CMRglucose should be considerably lower than 6:1. Obviously, most,

:CMRglucose measured or calculated should approach 6 [23]. The conclu-

:CMRlactate should not exceed

ing neural tissue activation, the expected ratio CMR <sup>O</sup> <sup>2</sup>

**40**

the ratio CMR <sup>O</sup> <sup>2</sup>

mitochondrial TCA cycle.

ratio CMR <sup>O</sup> <sup>2</sup>

**Figure 5.**

*Figure 3 and [36].*

Hyder et al. [23] used BOLD fMRI and the measurements used by Fox et al. [21] were even more cumbersome, involving the use of [15O]H2O, [15O]O2 and [15O]CO2.

While BOLD fMRI estimates yielded a CMRO2 :CMRglucose ratio value of 6:1, the 15O measurements produced a ratio value of 0.4:1. These completely opposing outcomes make one wonder whether or not the measurements performed using these two methods and the calculated values of CMRO2 they produced truly reflect the changes in the consumption of molecular oxygen upon neural activation. Could the direct measurements of CMRO2 , CMRglucose and CMRlactate done by Hu and Wilson [41] along with the indirect ones made by Fox et al. [21] and Hyder et al. [23], be reconciled such that a better picture of cerebral metabolic rates of activated neural tissue can be visualized? It is widely agreed that over 90% of the normal brain's energy production originates from glucose oxidation [21, 51]. The normal glucose concentration in the brain is ~2 mM and its normal lactate concentration is about half of that of glucose. Thus, it is safe to postulate that the normal resting brain is supplied with ample amounts of oxygen to continuously oxidize more than 90% of the brain glucose. However, glucose supplies to the normal brain are limited (only 40% of normal blood glucose level). Consequently, the increased rate of CBF along with the increased consumption of glucose upon activation [21, 23, 52] should supply all the oxygen necessary to match the increased demand, in contrast to the limited supplies of glucose. Low resolution techniques for the measurements of oxygen concentrations are unable to detect local fluctuations accurately if at all, which could explain why Fox et al. [21] reached the conclusion regarding the very low oxygen consumption during neural activation. Nonetheless, their conclusion that the energy demands of activated neural tissue are being met through glycolytic ATP production is most likely incorrect. In other words, undetectable or slightly detectable dip in tissue oxygen level upon activation is not necessarily an indication that oxygen is not consumed. The higher resolution of oxygen measurement afforded by polarography exemplifies the fact that local oxygen levels dipped upon stimulation and overshot upon its cessation ([41]; **Figures 3** and **4**). Although local fluctuations in tissue oxygen levels were evident, its overall tissue concentration did not significantly change and may even have risen somewhat above its baseline level. In contrast, both glucose and lactate levels were changed significantly from their baseline levels [24, 36, 41] (**Figures 3**–**5**). The fluctuations between lactate and oxygen were highly synchronized, indicating that lactate is being oxidized upon tissue activation. During the 20 min following the 10th stimulation, the tissue level of both oxygen and glucose climbed above the baseline level, while the high level of lactate gradually declined ([41]; **Figure 3**). These shifts seem to indicate that upon cessation of stimulation, as the tissue is recovering from activation and high energy demands, lactate becomes the preferred oxidative energy substrate, sparing glucose. That the cerebral tissue would prefer lactate over glucose, especially when the former is abundant, is reasonable, considering the fact that lactate oxidative mitochondria. Consequently, the increased rate of CBF along with the increased consumption of glucose upon activation [21, 23, 52] should supply all the oxygen necessary to match the increased demand, in contrast to the limited supplies of glucose. Low resolution techniques for the measurements of oxygen concentrations are unable to detect local fluctuations accurately if at all, which could explain why Fox et al. [21] reached the conclusion regarding the very low oxygen consumption during neural activation. Nonetheless, their conclusion that the energy demands of activated neural tissue are being met through glycolytic ATP production is most likely incorrect. In other words, undetectable or slightly detectable dip in tissue oxygen level upon activation is not necessarily an indication that oxygen is not consumed. The higher resolution of oxygen measurement afforded by polarography exemplifies the fact that local oxygen levels dipped upon stimulation and overshot upon its cessation (**Figures 3** and **4** and [41]). Although local fluctuations in tissue

oxygen levels were evident, its overall tissue concentration did not significantly change and may even have risen somewhat above its baseline level. In contrast, both glucose and lactate levels were changed significantly from their baseline levels (**Figures 3**–**5** and [24, 36, 41]). The fluctuations between lactate and oxygen were highly synchronized, indicating that lactate is being oxidized upon tissue activation. During the 20 min following the 10th stimulation, the tissue level of both oxygen and glucose climbed above the baseline level, while the high level of lactate gradually declined (**Figure 3** and [41]). These shifts seem to indicate that upon cessation of stimulation, as the tissue is recovering from activation and high energy demands, lactate becomes the preferred oxidative energy substrate, sparing glucose. That the cerebral tissue would prefer lactate over glucose, especially when the former is abundant, is reasonable, considering the fact that lactate oxidative utilization, in contrast to glucose, does not involve ATP investment prior to its utilization by mitochondria.

#### **4. CMRs measurements and their possible implications in brain disorders**

Energy metabolic interruptions are at the basis of several brain disorders and measuring CMRs of patients inflicted by such brain disorders can offer a potentially better diagnosis and treatment. Measurement of CMRO2 and CMRglucose have been performed regularly in numerous studies of cerebral ischemia in an effort to better understand the mechanisms of neuronal ischemic damage. CMRglucose measurement has been used in studying obsessive-compulsive disorder, mood disorder and depression, where the main aim is to follow changes in glucose metabolism in specific brain regions believed to be involved in these disorders. Other brain disorders where glucose metabolic rate has been measured in include amyotrophic lateral sclerosis, Alzheimer's disease, epilepsy, Parkinson's disease and Huntington's disease. The purpose behind the measurement of glucose cerebral metabolic rate when investigating diseases and disorders is usually to identify brain regions that are involved in a given disorder or disease, not to investigate how energy metabolism is being affected by the disease or the disorder. Also, the energy metabolic rates of brain tumors have received great attention due to the unique energy requirements of these tumors. Nevertheless, cerebral ischemia and traumatic brain injury (TBI) are the two disorders for which measurements of CMRO2 and CMRglucose are most abundant. The results of these measurements prompted proposals both for treatments and mechanisms of neuronal damage due to these insults. The most heralded hypothesis attempting to explain delayed neuronal cerebral ischemic damage [20] known as the lactic acidosis hypothesis, postulated the accumulation of lactic acid as the cause of that damage. Consequently, physicians dealing with stroke patients were encouraged to control blood glucose levels in these patients, based on the assumption that the higher the glucose level during cerebral ischemia, the higher the level of lactic acid produced and the damage it causes. The lactic acidosis hypothesis was discarded, although the practice of controlling the blood glucose level of stroke patients remained. To this end, lactate was shown to support neuronal recovery post-ischemia *in vitro* [53–55]. Moreover, higher glucose level pre-ischemia (hypoxia) appear to improve neuronal recovery post-ischemia *in vitro* [56] and any exacerbation of neuronal damage due to pre-ischemic hyperglycemia was shown to be induced by increased levels of stress hormone [57]. Similarly, experimental [58] and clinical studies [59–66] over the past two decades indicate that lactate supplementation after TBI improves post-injury outcome. Measurement of CMRlactate could greatly enhance our understanding of the role that this monocarboxylate plays in these two brain disorders.

**43**

**Author details**

Avital Schurr

provided the original work is properly cited.

Louisville, School of Medicine, Louisville, KY, USA

\*Address all correspondence to: avital.schurr@gmail.com

*Cerebral Energy Metabolism: Measuring and Understanding Its Rate*

A paradigm shift of a scientific model should, naturally, result in reconsideration of hypotheses and concepts that have been formulated on its foundation prior to its shift. The understanding of cerebral metabolic rates of energy substrates during rest and activation of neural tissue, the use of the method best suited for the measurement of these rates and the interpretation of the results have always relied on two fundamental assumptions. First, cerebral energy metabolism includes the obligatory glycolytic breakdown of glucose to pyruvate and the utilization of the latter by the mitochondrial TCA cycle and the electron transport chain with oxygen as its final receptor. Second, the activation of cerebral tissue is sustained by an increase in ATP production and therefore an increase in the consumption of glucose and oxygen. Two seminal papers that were published almost simultaneously [16, 21] have forced biochemists, and especially neuroscientists, to reassess these two basic postulates. The paper by Fox et al. [21] has perplexed many with its conclusion that the energy requirements of activated neural tissue are minimal and can be fulfilled by the glycolytic pathway alone (glucose → lactate + 2ATP). The paper by Schurr et al. [16] provoked great skepticism upon demonstrating that neural tissue can function and be activated when lactate is its sole oxidative energy substrate (lactate + 3O2 + mitochondria → pyruvate → TCA cycle → 3CO2 + 3H2O + 17ATP). While the proposal that lactate is a suitable oxidative energy substrate had faced strong skepticism for many years, it has gained greater support over the past three decades. The proposal that glycolysis could be served as the sole supplier of energy for the activated neural tissue still divides scientists working in this field. By accepting the proposed paradigm shift of glycolysis [24] and its application in the interpretation of the results obtained by Fox et al. [21], Hyder et al. [23, 67], Hu and Wilson [41] and many others, a scenario can be drawn where lactate is supplementing most if not all the energy requirements of activated neural tissue. The data and the line of reasoning presented here strongly argue against the conclusion that these energy requirements are solely fulfilled by glycolysis. Future studies of activated cerebral

*DOI: http://dx.doi.org/10.5772/intechopen.84376*

**5. Conclusions**

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium,

Department of Anesthesiology and Perioperative Medicine, University of

metabolic rates should include, along with the measurements of CMRO2

the measurement of CMRlactate. Resolving the existing debated issues of cerebral energy metabolism is paramount for our better understanding the many brain diseases and disorders. Hopefully, this chapter provides a possible resolution of some of these issues.

and CMRglucose,

### **5. Conclusions**

*Cellular Metabolism and Related Disorders*

**disorders**

oxygen levels were evident, its overall tissue concentration did not significantly change and may even have risen somewhat above its baseline level. In contrast, both glucose and lactate levels were changed significantly from their baseline levels (**Figures 3**–**5** and [24, 36, 41]). The fluctuations between lactate and oxygen were highly synchronized, indicating that lactate is being oxidized upon tissue activation. During the 20 min following the 10th stimulation, the tissue level of both oxygen and glucose climbed above the baseline level, while the high level of lactate gradually declined (**Figure 3** and [41]). These shifts seem to indicate that upon cessation of stimulation, as the tissue is recovering from activation and high energy demands, lactate becomes the preferred oxidative energy substrate, sparing glucose. That the cerebral tissue would prefer lactate over glucose, especially when the former is abundant, is reasonable, considering the fact that lactate oxidative utilization, in contrast to glucose, does not involve ATP investment prior to its utilization by mitochondria.

**4. CMRs measurements and their possible implications in brain** 

better diagnosis and treatment. Measurement of CMRO2

(TBI) are the two disorders for which measurements of CMRO2

boxylate plays in these two brain disorders.

Energy metabolic interruptions are at the basis of several brain disorders and measuring CMRs of patients inflicted by such brain disorders can offer a potentially

performed regularly in numerous studies of cerebral ischemia in an effort to better understand the mechanisms of neuronal ischemic damage. CMRglucose measurement has been used in studying obsessive-compulsive disorder, mood disorder and depression, where the main aim is to follow changes in glucose metabolism in specific brain regions believed to be involved in these disorders. Other brain disorders where glucose metabolic rate has been measured in include amyotrophic lateral sclerosis, Alzheimer's disease, epilepsy, Parkinson's disease and Huntington's disease. The purpose behind the measurement of glucose cerebral metabolic rate when investigating diseases and disorders is usually to identify brain regions that are involved in a given disorder or disease, not to investigate how energy metabolism is being affected by the disease or the disorder. Also, the energy metabolic rates of brain tumors have received great attention due to the unique energy requirements of these tumors. Nevertheless, cerebral ischemia and traumatic brain injury

most abundant. The results of these measurements prompted proposals both for treatments and mechanisms of neuronal damage due to these insults. The most heralded hypothesis attempting to explain delayed neuronal cerebral ischemic damage [20] known as the lactic acidosis hypothesis, postulated the accumulation of lactic acid as the cause of that damage. Consequently, physicians dealing with stroke patients were encouraged to control blood glucose levels in these patients, based on the assumption that the higher the glucose level during cerebral ischemia, the higher the level of lactic acid produced and the damage it causes. The lactic acidosis hypothesis was discarded, although the practice of controlling the blood glucose level of stroke patients remained. To this end, lactate was shown to support neuronal recovery post-ischemia *in vitro* [53–55]. Moreover, higher glucose level pre-ischemia (hypoxia) appear to improve neuronal recovery post-ischemia *in vitro* [56] and any exacerbation of neuronal damage due to pre-ischemic hyperglycemia was shown to be induced by increased levels of stress hormone [57]. Similarly, experimental [58] and clinical studies [59–66] over the past two decades indicate that lactate supplementation after TBI improves post-injury outcome. Measurement of CMRlactate could greatly enhance our understanding of the role that this monocar-

and CMRglucose have been

and CMRglucose are

**42**

A paradigm shift of a scientific model should, naturally, result in reconsideration of hypotheses and concepts that have been formulated on its foundation prior to its shift. The understanding of cerebral metabolic rates of energy substrates during rest and activation of neural tissue, the use of the method best suited for the measurement of these rates and the interpretation of the results have always relied on two fundamental assumptions. First, cerebral energy metabolism includes the obligatory glycolytic breakdown of glucose to pyruvate and the utilization of the latter by the mitochondrial TCA cycle and the electron transport chain with oxygen as its final receptor. Second, the activation of cerebral tissue is sustained by an increase in ATP production and therefore an increase in the consumption of glucose and oxygen. Two seminal papers that were published almost simultaneously [16, 21] have forced biochemists, and especially neuroscientists, to reassess these two basic postulates. The paper by Fox et al. [21] has perplexed many with its conclusion that the energy requirements of activated neural tissue are minimal and can be fulfilled by the glycolytic pathway alone (glucose → lactate + 2ATP). The paper by Schurr et al. [16] provoked great skepticism upon demonstrating that neural tissue can function and be activated when lactate is its sole oxidative energy substrate (lactate + 3O2 + mitochondria → pyruvate → TCA cycle → 3CO2 + 3H2O + 17ATP). While the proposal that lactate is a suitable oxidative energy substrate had faced strong skepticism for many years, it has gained greater support over the past three decades. The proposal that glycolysis could be served as the sole supplier of energy for the activated neural tissue still divides scientists working in this field. By accepting the proposed paradigm shift of glycolysis [24] and its application in the interpretation of the results obtained by Fox et al. [21], Hyder et al. [23, 67], Hu and Wilson [41] and many others, a scenario can be drawn where lactate is supplementing most if not all the energy requirements of activated neural tissue. The data and the line of reasoning presented here strongly argue against the conclusion that these energy requirements are solely fulfilled by glycolysis. Future studies of activated cerebral metabolic rates should include, along with the measurements of CMRO2 and CMRglucose, the measurement of CMRlactate. Resolving the existing debated issues of cerebral energy metabolism is paramount for our better understanding the many brain diseases and disorders. Hopefully, this chapter provides a possible resolution of some of these issues.

### **Author details**

Avital Schurr Department of Anesthesiology and Perioperative Medicine, University of Louisville, School of Medicine, Louisville, KY, USA

\*Address all correspondence to: avital.schurr@gmail.com

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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#### *Cerebral Energy Metabolism: Measuring and Understanding Its Rate DOI: http://dx.doi.org/10.5772/intechopen.84376*

(Chap. 31). In: Siegle GJ, Agranoff BW, Albers RW, Molinoff PB, editors. Basic Neurochemistry. 5th ed. New York, NY: Raven Press; 1994. pp. 645-680

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*Cellular Metabolism and Related Disorders*

The Journal of Biological Chemistry. 2013;**288**:25309-25317. DOI: 10.1074/jbc. oxygenation. Proceedings of the National Academy of Sciences of the United States of America. 1990;**87**:9868-9872. DOI:

[40] Bentley WJ. Oxygen Polarography in the Awake Macaque: Bridging BOLD fMRI and Electrophysiology. 2014. Available from: http://openscholarship.

[41] Hu Y, Wilson GS. A temporary local energy pool coupled to neuronal activity: fluctuations of extracellular lactate levels in rat brain monitored with rapid-response enzyme-based sensor. Journal of Neurochemistry.

10.1046/j.1471-4159.1997.69041484.x

[42] Pellerin L, Magistretti PJ. Food for thought: Challenging the dogmas. Journal of Cerebral Blood Flow and Metabolism.

2003;**23**:1282-1286. DOI: 10.1097/01.

[43] Kasischke KA, Vishwasrao HD, Fisher PJ, Zipfel WR, Webb WW. Neural activity triggers neuronal oxidative metabolism followed by astrocytic glycolysis. Science. 2004;**305**:99-103.

[44] Aubert A, Costalat R, Magistretti PJ, Pellerin L. Brain lactate kinetics: Modeling evidence for neuronal lactate uptake upon activation. Proceedings of the National Academy of Sciences of the United States of America. 2005;**102**:16448-16453. DOI: 10.1073/

[45] Medina JM, Tabernero A. Lactate utilization by brain cells and its role in CNS development. Journal of Neuroscience Research. 2005;**179**:2-10.

[46] Serres S, Bezancon E, Franconi J-M, Merle M. Ex vivo NMR study of lactate metabolism in rat brain under various depressed states. Journal of

WCB.0000096064.12129.3D

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pnas.0505427102

DOI: 10.1002/jnr.20336

10.1073/pnas.87.24.9868

wustl.edu/etd/1218

1997;**69**:1484-1490. DOI:

[33] Brooks GA, Dubouchaud H, Brown M, Sicurello JP, Butz CE. Role of mitochondrial lactate dehydrogenase and lactate oxidation in the intracellular lactate shuttle. Proceedings of the National Academy of Sciences of the United States of America. 1999;**96**: 1129-1134. DOI: 10.1073/pnas.96.3.1129

[34] Havel RJ, Watkins E, Gullixson KS. The metabolism of lactate and pyruvate in children with congenital heart disease. Circulation. 1950;**2**:536-544

[35] Schurr A, Payne RS. Lactate, not pyruvate, is neuronal aerobic glycolysis end product: An in vitro

Neuroscience. 2007;**147**:613-619. DOI: 10.1016/j.neuroscience.2007.05.002

[37] Sokoloff L, Reivich M, Kennedy C, Des Rosiers MH, Patlak CS, Pettigrew KD, et al. The [14C]deoxyglucose method for the measurement of local cerebral glucose utilization: Theory, procedure, and normal values in the conscious and anesthetized albino rat. Journal of Neurochemistry. 1977;**28**: 897-916. DOI: 10.1111/j.1471-4159.1977.

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electrophysiological study.

fphar.2011.00096

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M113.476648

**46**

tb10649.x

[48] Fillenz M. The role of lactate in brain metabolism. Neurochemistry International. 2005;**47**:413-417. DOI: 10.1016/j.neuint.2005.05.011

[49] Korf J. Is brain lactate metabolized immediately after neuronal activity through the oxidative pathway? Journal of Cerebral Blood Flow and Metabolism. 2006;**26**:1584-1586. DOI: 10.1038/ sj.jcbfm.9600321

[50] Chambers TW, Daly TP, Hockley A, Brown AM. Contribution of glycogen in supporting axon conduction in the peripheral and central nervous systems: The role of lactate. Frontiers in Neuroscience. 2014;**8**:378. DOI: 10.3389/ fnins.2014.00378

[51] Siesjo BK. Brain Energy Metabolism. New York: John Wiley and Sons; 1978

[52] Ueki M, Linn F, Hossmann K-A. Functional activation of cerebral blood flow and metabolism before and after global ischemia of rat brain. Journal of Cerebral Blood Flow and Metabolism. 1988;**8**:486-494

[53] Schurr A, Dong W-Q , Reid KH, West CA, Rigor BM. Lactic acidosis and recovery of neuronal function following cerebral hypoxia in vitro. Brain Research. 1988:311-314

[54] Schurr A, Rigor BM. Brain anaerobic lactate production: A suicide note or a survival kit? Developmental Neuroscience. 1998;**20**:348-357. DOI: 10.1159/000017330

[55] Schurr A, Payne RS, Miller JJ, Tseng MT, Rigor BM. Blockade of lactate transport exacerbates delayed neuronal damage in a rat model of cerebral ischemia. Brain Research. 2001;**895**:268-272

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[58] Rice AC, Zsoldos R, Chen T, Wilson MS, Alessandri B, Hamm RJ, et al. Lactate administration attenuates cognitive deficits following traumatic brain injury. Brain Research. 2002;**928**:156-159. DOI: 10.1016/ S0006-8993(01)03299-1

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[60] Ichai C, Armando G, Orban JC, Berthier F, Rami L, Samat-Long C, et al. Sodium lactate versus mannitol in the treatment of intracranial hypertensive episodes in severe traumatic braininjured patients. Intensive Care Medicine. 2009;**35**:471-479. DOI: 10.1007/s00134-008-1283-5

[61] Ichai C, Payen JF, Orban JC, Quintard H, Roth H, Legrand R, et al. Half-molar sodium lactate infusion to prevent intracranial hypertensive episodes in severe traumatic brain injured patients: A randomized controlled trial. Intensive Care Medicine. 2013;**39**:1413-1422. DOI: 10.1007/s00134-013-2978-9

[62] Jalloh I, Helmy A, Shannon RJ, Gallagher CN, Menon DK, Carpenter KL, et al. Lactate uptake by the injured human brain: Evidence from an arteriovenous gradient and cerebral microdialysis study. Journal of Neurotrauma. 2013;**30**:2031-2037. DOI: 10.1089/neu.2013.2947

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**49**

Section 3

Diabetes Mellitus
