3.2 Choline and Its Products
R. J. Wurtman .M. Cansev .I. H. Ulus
1 Introduction ...................................................................................... 3
2 Choline in the Blood . ............................................................................ 3
2.1 Sources of Plasma Choline . . . . . . . . ................................................................. 4
2.1.1 Dietary Choline ..................................................................................... 4
2.1.2 Endogenous Choline Synthesis . . . . ................................................................. 5
2.1.3 Choline‐Containing Membrane Phospholipids . . . . . . . . . . . . . . . . . ................................... 9
2.2 Fates of Circulating Choline . . . . . . . ................................................................ 10
2.2.1 Choline as a Source of Methyl Groups (Choline Oxidase System) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12
2.3 Effects of Physiologic or Pathologic States on Plasma Choline .................................. 15
3 Choline in the Brain . ........................................................................... 16
3.1 Sources of Brain Choline . . . . . . . . . . ................................................................ 16
3.1.1 Uptake of Circulating Choline into the Brain . . . . . . . . . . . . . . . . . . .................................. 16
3.1.2 Liberation from Membrane PC . . . . ................................................................ 17
3.1.3 Reutilization of Choline Formed from Hydrolysis of Acetylcholine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18
3.1.4 De Novo Synthesis of Phosphatidylcholine and Choline . . . . . . . .................................. 18
4 Brain Proteins that Interact with Choline ..................................................... 18
4.1 Enzymes . . . . . . . . .................................................................................... 19
4.1.1 Choline Acetyltransferase . . . . . . . . . . ................................................................ 19
4.1.2 Choline Kinase . .................................................................................... 20
4.2 Transport Proteins . . . . . . . . . . . . . . . . . ................................................................ 20
4.2.1 Facilitated‐Diffusion Carrier at Blood–Brain Barrier . . . . . . . . . . . .................................. 20
4.2.2 Choroid Plexus Choline Transporter . . . . . . . . ...................................................... 21
4.2.3 High‐Affinity Uptake Protein in Cholinergic Terminals . . . . . . . . .................................. 21
4.2.4 Low‐Affinity Cellular Uptake Protein . . . . . . . ...................................................... 22
4.3 Receptors . . . . . . . .................................................................................... 22
5 Utilization of Choline in Brain ................................................................. 24
5.1 Biosynthesis of Acetylcholine . . . . . . ................................................................ 24
5.2 Biosynthesis of Choline‐Containing Phosphatides . . . . . . . . . . . . . . .................................. 25
5.2.1 CTP:Phosphocholine Cytidylyltransferase . . . ...................................................... 29
5.2.2 CDP‐Choline:1,2‐Diacylglycerol Cholinephosphotransferase . . . .................................. 30
5.2.3 Uptake of Uridine and Cytidine into Brain Cells . . . . . . . . . . . . . . . .................................. 31
5.2.4 Phosphorylation of Uridine and Cytidine to UTP and CTP . . . .................................. 31
#Springer-Verlag Berlin Heidelberg 2007
6 Physiological and Behavioral Effects of Choline ............................................... 32
6.1 Blood Pressure . .................................................................................... 32
6.2 Body Temperature . . . . . . . . .......................................................................... 33
6.3 Pain ................................................................................................. 33
6.4 Neuroendocrine Effects . . .......................................................................... 33
6.5 Peripheral Metabolism . . . .......................................................................... 33
6.6 Behavior . . . . . . . . .................................................................................... 34
6.7 Drug Interactions . . . . . . . . .......................................................................... 34
6.8 Neuroprotective and Cytoprotective Effects . . . . . . . . . . . ............................................ 35
7 Effects of Exogenous CDP‐Choline .............................................................. 35
7.1 Hypoxia and Ischemia . . . .......................................................................... 36
7.2 Head Trauma (Cranio‐Cervical Trauma) . . . . . . . . . . . . . . ............................................ 37
7.3 Induced Lesions .................................................................................... 37
7.4 Other Effects . . . .................................................................................... 37
7.5 Clinical Studies . .................................................................................... 38
8 Choline in Autonomic and Motor Neurons .................................................... 38
23.2 Choline and its products acetylcholine and phosphatidylcholine
Abstract: Choline, a quaternary amine obtained largely from the diet but also synthesized in the brain and,
especially, liver, is an essential precursor of the neurotransmitter acetylcholine (ACh) and of the major
membrane constituent phosphatidylcholine (PC). Plasma choline concentrations can vary over a ﬁvefold
range depending on the choline contents of the foods being digested. Since choline readily crosses the
blood–brain barrier (BBB) through an unsaturated facilitated‐diffusion system, these plasma changes can
produce parallel changes in brain choline levels. In addition, since the enzymes that convert choline to ACh
[choline acetyltransferase (ChAT)] and PC’s precursor phosphocholine [choline kinase (CK)] are also
poorly saturated with their choline substrate, increases in plasma choline can enhance the formation of ACh
and phosphocholine, and the release of ACh. The subsequent conversion of phosphocholine to PC is
increased if PC’s other circulating precursors (uridine and omega‐3 fatty acids) are provided. This leads
to an increase in the levels of synaptic membrane within the brain. Choline is principally metabolized in
the liver to betaine, which provides a source of methyl groups for the regeneration of methionine and
Choline (2‐hydroxyethyl‐trimethyl‐ammonium), a simple but unusual compound, consists of a 2‐carbon
chain in which one carbon is attached to a hydroxyl group and the other to an aminenitrogen (>Figure 3.2‐1).
The particularly unusual quality of choline is that this amine nitrogen bonds with a total of four
hydrogen or carbon atoms instead of with the usual three, and thus carries a partial positive charge.
Though choline is not an amino acid, it shares with that family of compounds the property of being
present in cells both in free form and—like the amino acids in proteins—within subunits (e.g., PC) of a
macromolecule (biologic membranes). Moreover, like tyrosine, tryptophan (Cansev and Wurtman,
2006), and histidine, choline is the precursor of a neurotransmitter (ACh), and also like tyrosine and
histidine, choline must be obtained from both endogenous synthesis and dietary sources.
This chapter describes the choline in the blood and, particularly, in the brain, —its sources; its uses to
make membrane phospholipids and ACh; and its other biologic effects.
2 Choline in the Blood
Choline is a normal constituent of the plasma (>Table 3.2‐1), present as the free base (Cohen and
Wurtman, 1975; Hirsch et al., 1978; Savendahl et al., 1997); as a constituent of phospholipids [including
PC; sphingomyelin (SM); lyso‐PC; choline‐containing plasmalogens; and the platelet‐activating factor
(PAF)] and as PC’s water‐soluble metabolites [principally phosphocholine and glycerophosphocholine
(GPC) (Hirsch et al., 1978; Klein et al., 1993; Ilcol et al., 2005a)]. Free choline is also found in other
biological ﬂuids (>Table 3.2‐1). In blood, choline is concentrated within erythrocytes (10–150 mM;
Structure of choline
Choline and its products acetylcholine and phosphatidylcholine 3.2 3
Jope et al., 1980; Stoll et al., 1991) through the action of an uptake molecule that is unsaturated (K
mM; Riley et al., 1997) at normal plasma choline concentrations.
2.1 Sources of Plasma Choline
Plasma choline derives from three sources—dietary choline, consumed as the free base or as a constituent of
phospholipid molecules; endogenous synthesis, principally in liver; and liberation from its reservoir within
the membrane phosphatides of all mammalian cells.
2.1.1 Dietary Choline
Choline is present within many foods (>Table 3.2‐2), principally as the free molecule or as phosphatides,
and its plasma levels can rapidly increase severalfold after ingestion of choline‐rich foods (Hirsch et al.,
1978). For example, the consumption of a 5‐egg omelet (containing about 1.3 g of choline) by humans
increased these levels from 9.8 mM to 36.6 mM within 4 h (Hirsch et al., 1978). Prolonged fasting reduced
human plasma choline levels from 9.5 mM to 7.8 mM after 7 days (Savendahl et al., 1997). Similarly, removal
of all choline‐containing foods from the diet for 17–19 days gradually lowered plasma choline, from 10.6 mM
to 8.4 mM in humans (Zeisel et al., 1991) and from 12.1 mM to 6.3 mM in rats (Klein et al., 1998), indicating
that plasma choline can be partially but not fully sustained by release from endogenous stores.
Dietary PC is deacylated within the gut to form lyso‐PC. About half of this product is further degraded
to free choline within the gut or liver. The remainder is reacylated to regenerate PC (Houtsmuller, 1979),
which is then absorbed into the lymphatic circulation (Fox et al., 1979) and eventually enters the blood
Much of the dietary choline that reaches the liver through the portal circulation is destroyed by
oxidation to betaine, as described later (>Figure 3.2‐2), ultimately providing methyl groups that can be
used to regenerate S‐adenosylmethionine (SAM) from homocysteine. The rest of the choline in portal
venous blood passes into the systemic circulation (Fox et al., 1979; Houtsmuller, 1979).
In 1998, the Food and Nutrition Board (FNB) of the US Institute of Medicine established a dietary
reference intake (DRI) for choline. Since the FNB did not believe that existing scientiﬁc evidence allowed
calculation of a recommended daily allowance (RDA) for choline, it instead set an adequate (daily) intake
level (AI), and an upper (daily) limit (UL) that should not be exceeded (>Table 3.2‐3). The main criteria
for determining the AI and UL were, respectively, the amount of choline needed to prevent liver damage,
Free choline concentrations in human body ﬂuids
Body ﬂuids Free choline (mM) References
Plasma 7.0–13.0 Holm et al. (2003)
Serum 9.0–13.3 Holm et al. (2003); Ilcol et al. (2002b, 2005a)
Urine 15.5 8.8 Buchman et al. (1999)
Cerebrospinal ﬂuid 0.7–2.5 Flentge et al. (1984); Ikeda et al. (1990); Toghi et al. (1996)
Amniotic ﬂuid 22.8–24.5 Ilcol et al. (2002e)
Colostrum 132 21 Ilcol et al. (2005a)
Breast milk 110–300 Holmes et al. (2000); Ilcol et al. (2005a)
Semen 17,000–24,000 Takatori et al. (1984); Manabe et al. (1991)
Peritoneal dialysate 14–28 Hjelle et al. (1993); Ilcol et al. (2002b)
Note: Values are given as the range of the means from the cited references, or as the mean SEM
43.2 Choline and its products acetylcholine and phosphatidylcholine
and the choline intake associated with choline’s most sensitive adverse effect, i.e., hypotension (see Dietary
Reference Intakes, Institute of Medicine, National Academy of Sciences USA, 1998). Subsequent studies
have shown that the enzymes (described later) that synthesize and metabolize choline can be affected by
common genetic polymorphisms, which cause important person‐to‐person variations in dietary choline
needs (da Costa et al., 2006). For further details about dietary reference intakes and the choline contents
of various foods, the reader is referred to the ofﬁcial websites of the Institute of Medicine (http://www.
nap.edu/catalog/6015.html#toc) and the USDA (http://www.nal.usda.gov/fnic/foodcomp/Data/Choline/
2.1.2 Endogenous Choline Synthesis
Endogenous choline is produced, principally in liver (Bremer and Greenberg, 1960) but also to a small
extent within brain (Blusztajn et al., 1979; Crews et al., 1980), by the sequential addition of three methyl
groups to the amine nitrogen of phosphatidylethanolamine (PE); this forms PC, which can then be broken
down to liberate the choline (>Figure 3.2‐3). The methylation reactions are catalyzed by two enzymes,
phosphatidylethanolamine‐N‐methyltransferase (PEMT1; EC: 18.104.22.168), which converts PE to its mono-
methyl derivative, and phosphatidyl‐N‐methylethanolamine‐N‐methyltransferase (PEMT2; EC: 22.214.171.124),
which adds the second and third methyl groups (A single enzyme may catalyze all three methylations in
liver). Both enzymes use SAM as the methyl donor (Bremer and Greenberg, 1960; Hirata et al., 1978). Their
s for SAM are 2–4 10
M and 20–110 10
M, respectively (Crews et al., 1980; Blusztajn et al.,
1982; Hitzemann, 1982; Percy et al., 1982), whereas brain SAM concentrations are 10–17 mg/g wet weight
[50–85 mM assuming about 50% of the brain mass is aqueous (Wurtman and Rose, 1970; Ordonez and
Wurtman, 1974)]. Hence, PEMT1 is probably fully saturated with SAM whereas PEMT2 is not.
Choline contents of common foods
Food Choline GPC PCho PC SM Total Choline
Egg yolk 1 1 1 634 45 682
Beef liver 61 83 11 245 24 424
Chicken liver 69 5 6 213 14 307
Cereals, ready to eat or wheat germ 69 33 4 44 0 150
Pork or bacon 12 18 3 86 10 129
Cake, chocolate, without frosting 5 61 1 58 2 127
Coffee, instant, decaffeinated 93 8 0 0 0 101
Cauliﬂower 24.5 0.7 1.8 12.1 0 39.1
Food Choline GPC PCho PC SM Total Choline
Olive oil 0 0.3 0 0 0 0.3
Kale 0.1 0 0 0.3 0 0.4
Iced tea 0.4 0 0 0 0 0.4
Egg white 0.2 0.6 0 0.3 0 1.1
Apple juice 0.7 0.7 0 0.4 0 1.8
Coffee, brewed from grounds 1.9 0.7 0 0.4 0 2.6
Note: Data are given as mg choline moiety/100 g of food. Foods are grouped based on having relatively high or low choline
contents. Data from USDA Database for the Choline Content of Common Foods, 2004
GPC, glycerophosphocholine; PCho, phosphocholine; PC, phosphatidylcholine; SM, sphingomyelin
Choline and its products acetylcholine and phosphatidylcholine 3.2 5
Metabolism of choline to betaine, methionine and, S‐adenosylmethionine (SAM). THF, Tetrahydrofolate;
5,10‐MTHF, 5,10‐methylene‐tetrahydrofolate; 5‐methyl‐THF, 5‐methyl‐tetrahydrofolate; B12, Vitamin B12;
SAM, S‐adenosylmethionine; SAH, S‐adenosylhomocysteine
63.2 Choline and its products acetylcholine and phosphatidylcholine
The gene for human PEMT2 has been localized to chromosome 17p11.2 (Walkey et al., 1999), and
cDNA for PEMT2 has been cloned from rat liver and expressed in COS‐1 cells (Cui et al., 1993). PEMT
activity was identiﬁed in membranous fractions from homogenates of rat and bovine brain (Blusztajn et al.,
1979; Mozzi and Porcellati, 1979; Crews et al., 1980); highest speciﬁc activies were present in synaptosomes
(Blusztajn et al., 1979; Crews et al., 1980) suggesting that nerve terminals are able to synthesize choline. In
the course of these transmethylations, the phosphatide intermediates ‘‘ﬂip’’ from the membrane’s cytoplas-
mic side, where most of the less‐polar PE and phosphatidylserine (PS) are found, to the more polar external
leaf (Hirata and Axelrod, 1978). PE itself can be formed in liver, kidney, or brain from free ethanolamine,
via the CDP‐ethanolamine cycle (or ‘‘Kennedy Cycle’’; >Figure 3.2‐4) described later, or from the
decarboxylation of PS (Kennedy, 1956; Borkenhagen et al., 1961). PS is produced, in nerve terminals
(Holbrook and Wurtman, 1988) and elsewhere, by the process of ‘‘base‐exchange,’’ in which a serine
molecule substitutes for the ethanolamine in PE or the choline in PC.
Free choline is liberated from newly synthesized PC and from PC molecules formed from preexisting
choline, by a family of enzymes, the phospholipases (>Figure 3.2‐5). Phospholipase D (PLD) acts directly
on the choline/phosphate bond of PC to generate choline and phosphatidic acid (>Figure 3.2‐5a).
Phospholipase A2 (PLA2) acts on the bond connecting a fatty acid to the hydroxyl group on PC’s 2‐carbon
to yield that fatty acid [often arachidonic acid (AA) or docosahexaenoic acid (DHA)] and lyso‐PC
(>Figure 3.2‐5b). This lyso‐PC can then be further metabolized to choline, either directly, through the
action of a phosphodiesterase, or ﬁrst to GPC, by phospholipase A1 (PLA1), and then to choline by a
Proposed adequate (i.e., minimum) daily choline intake, and upper limit (which should not be exceeded)
A. Adequate Intakes (AI)
Age Gender AI (mg/day)
0–6 months Both 125
7–12 months Both 150
1–3 years Both 200
4–8 years Both 250
9–13 years Male 375
14–18 years Male 550
9–13 years Female 375
14–18 years Female 400
Above 19 years Male 550
Above 19 years Female 425
B. Upper Allowable Intakes (UL, Upper Limit)
Life stage Age UL (g/day)
Infancy 0–12 months N/A
Childhood 1–8 years 1
9–13 years 2
Adolescence 14–18 years 3
Pregnancy 14–18 years 3
19 years and older 3.5
Lactation 14–18 years 3
19 years and older 3.5
Not possible to establish; sources of intake should only be mother’s milk and infant formulas
Choline and its products acetylcholine and phosphatidylcholine 3.2 7
Sequential methylation of phosphatidylethanolamine (PE) to phosphatidylcholine (PC). PEMT‐I, Phosphatidyl-
ethanolamine‐N‐methyltransferase; PEMT‐II, phosphatidyl‐N‐methylethanolamine‐N‐methyltransferase; SAM,
S‐adenosylmethionine; SAH, S‐adenosylhomocysteine
83.2 Choline and its products acetylcholine and phosphatidylcholine
phosphatase. Phospholipase C (PLC) acts on the bond connecting the phosphate and the hydroxyl group on
PC’s 3‐carbon to yield diacylglycerol (DAG) and phosphocholine; the phosphocholine can then be
metabolized to free choline through the action of a phosphatase (>Figure 3.2‐5c).
It is estimated that, on average, about 15% of the free choline that enters the human blood stream
derives from endogenous synthesis, the rest coming principally from dietary sources (Zeisel, 1981). Acute
or chronic liver disease or deﬁciencies in methionine, folic acid, or vitamin B12 intake could thus lower
plasma choline levels by impairing hepatic PC synthesis (>Table 3.2‐4).
2.1.3 Choline‐Containing Membrane Phospholipids
Cellular membranes contain most of the choline in the body, principally in the form of the phosphatide PC,
but also as PC’s products SM (>Figure 3.2‐6) and lyso‐PC (>Figure 3.2‐5b), or as less‐abundant choline‐
containing phospholipids like the PAF (1‐O‐alkyl‐2‐acetyl‐sn‐glycero‐3‐phosphocholine). Membranes also
contain the phosphatides PS, PE, and phosphatidylinositol (PI), as well as speciﬁc proteins, cholesterol,
and various minor lipids. The quantities of choline present in brain as PC (2–2.5 mmoles/g) or as SM
(0.25 mmoles/g) are orders of magnitude greater than those of free choline (30–60 mM). The proportion
of any membrane’s phospholipids represented by PC can vary depending on the species and age of the
animal, the particular brain region or cell type being studied, and the membrane’s function within the cell
(e.g., nuclear membrane and plasma membrane) (Suzuki, 1981). In the gray matter of human brain, PC
constitutes 42% of total phospholipids and SM 10%; in white matter, the proportions of PC and SM are
33% and 15%, respectively (Suzuki, 1981).
Moreover, ‘‘PC’’ is highly heterogeneous, actually representing a family of compounds with differing
fatty acid compositions (>Figure 3.2‐3; Lee and Hajra, 1991) and, consequently, differing chemical and
Biosynthesis of phosphatidylcholine (PC) from preexisting choline through the CDP‐choline cycle (‘‘Kennedy
cycle’’). Synthesis of PC is shown here. Synthesis of phosphatidylethanolamine (PE) is similar except that it uses
ethanolamine instead of choline. Boxes indicate compounds most or all of which must be taken up into the
brain from the circulation. Uridine is the principal circulating precursor of the CTP (cytidine‐50‐triphosphate)
needed for PC and PE synthesis through the Kennedy pathway, and exogenous cytidine is rapidly deaminated
to uridine in humans (Wurtman et al., 2000). CK, Choline Kinase; CT: CTP: phosphocholine cytidylyltransferase;
CPT: CDP‐choline, 1,2‐diacylglycerol cholinephosphotransferase; PUFA, Polyunsaturated fatty acid; DHA, Doc-
osahexaenoic acid; EPA, Eicosapentaenoic acid; AA, Arachidonic acid
Choline and its products acetylcholine and phosphatidylcholine 3.2 9
physical properties. The fatty acid in the C‐1 position of PC tends most often to be saturated, e.g., stearic or
palmitic acid, whereas that in position C‐2 is more likely to be monounsaturated (oleic acid) or polyunsat-
urated [e.g., the omega‐3 fatty acids (DHA; 22:6) and eicosapentenoic acid (EPA; 20:5) or the omega‐6 fatty
acid (AA; 20:4)]. Newly synthesized phosphatide molecules contain relatively larger quantities of polyun-
saturated fatty acids (PUFA) than the phosphatide molecules present at steady state (Tacconi and Wurtman,
1985). This reﬂects either faster turnover of PUFA‐containing phosphatides, or their rapid deacylation
followed by reacylation with more saturated fatty acid species (Houtsmuller, 1979), or both. Membranes of
retinal and brain cells are especially rich in PUFA, particularly DHA [which comprises about 20% of the
total fatty acids in retinal phospholipids (Futterman and Andrews, 1964; Martinez, 1992) and about 7% of
those in brain phospholipids (Martinez, 1992), respectively]. As described later, administration of supple-
mental DHA accelerates PC synthesis and increases brain levels of PC and other phosphatides (Wurtman
et al., 2006).
2.2 Fates of Circulating Choline
Plasma choline can be taken up into the brain and other tissues for further metabolism, or, as discussed
later, oxidized—principally in liver and kidney—to form betaine, a source of methyl groups. This latter
process involves two enzymes, choline dehydrogenase and betaine dehydrogenase, usually collectively
termed ‘‘choline oxidase’’ (>Figure 3.2‐2). Negligible choline dehydrogenase activity (<1% of that found
in liver and kidney) was observed in rat brain in vitro (Haubrich et al., 1979) and no evidence exists that the
enzyme in brain actually converts choline to betaine in vivo; hence, we do not include the choline oxidase
system among the brain proteins, discussed later, that interact directly with choline. Humans (Buchman
Phospholipases that metabolize phosphatidylcholine (PC). (a)Phospholipase D; (b)Phospholipase A2;
(c) Phospholipase C. Boxes surrounding the portions of the PC molecule differentiate the glycerol, fatty acid
(R1 and R2), and choline moieties. Lyso‐PC, Lyso‐Phosphatidylcholine; LPCAT, Lyso‐Phosphatidylcholine acyl-
transferase; GPC‐PD, Glycerophosphocholine phosphodiesterase; GPC‐CPD, Glycerophosphocholine choline-
phosphodiesterase; DAG, Diacylglycerol
10 3.2 Choline and its products acetylcholine and phosphatidylcholine
.Figure 3.2‐5 (continued)
Choline and its products acetylcholine and phosphatidylcholine 3.2 11
et al., 1994) and laboratory animals (Acara and Rennick, 1973; Acara et al., 1979) also excrete small
amounts of unchanged choline into the urine (>Table 3.2‐1) by glomerular ﬁltration followed by partial
renal tubular reabsorbtion. Dietary choline or choline secreted into the gut can be broken down by
intestinal bacteria to form trimethylamine and related amine products (de la Huerga and Popper, 1951;
Neill et al., 1978). This process is responsible for the ‘‘ﬁshy odor’’ sometimes detected in people taking large
doses of choline supplements (Rehman, 1999).
2.2.1 Choline as a Source of Methyl Groups (Choline Oxidase System)
The choline oxidase system in mammals is composed of two enzymes; in microorganisms a single enzyme,
also termed ‘‘choline oxidase’’ (EC 126.96.36.199) converts choline to betaine. In mammals, the choline is ﬁrst
oxidized to betaine aldehyde by choline dehydrogenase (EC 188.8.131.52) (>Figure 3.2‐2), an enzyme located at
(or bound to) the inner membrane of mitochondria (Streumer‐Svobodova and Drahota, 1977; Lin and Wu,
1986). This enzyme can also convert the aldehyde to betaine; however, unlike the choline oxidase of
microorganisms, its afﬁnity for betaine aldehyde is very low (only about 5% its afﬁnity for choline), so
choline dehydrogenase has only a minor effect on net betaine synthesis (Tsuge et al., 1980). This enzyme is a
monomeric ﬂavoprotein with a molecular weight of 61.000 Da (Lin and Wu, 1986); its activity requires FAD
(Rothschild et al., 1954) and molecular oxygen serves as the primary electron acceptor (Zhang et al., 1992).
.Figure 3.2‐5 (continued)
12 3.2 Choline and its products acetylcholine and phosphatidylcholine
Effects of physiologic or pathologic conditions on circulating free choline concentrations in humans
(mM) Sample References
Preterm newborns (0–2 days) 21.2–33.3 S Buchman et al. (2001); Ilcol et al. (2005a)
Newborns at term (0–2 days) 28.1–68.1 S Zeisel et al. (1980a); Buchman et al. (2001); Ilcol
et al. (2005a)
At postnatal age 3–28 days 24.2–31.5 S Ilcol et al. (2005a)
At postnatal age 31–365 days 16.3–20.9 S Ilcol et al. (2005a)
At postnatal age366–730 days 12.8–16.3 P þS Buchman et al. (2001); Ilcol et al. (2005a)
Children (2–12 years old) 10.9–12.9 S Ilcol et al., 2005a
Adults (20–65 years old) 9.5–13.3 P þS Savendahl et al. (1997); Ilcol et al. (2002b, 2005a)
1‐week starved adults 7.5 0.3 P þS Savendahl et al. (1997)
Nonpregnant women 10.6 0.5 S Ilcol et al. (2002e)
Pregnant women 14.5–17.1 S Ulus et al. (1998); Ilcol et al. (2002e)
Nonlactating women 10.6 0.6 S Ilcol et al. (2005a)
14.7–19.2 S Ilcol et al. (2005a)
Before childbirth 15.4–16.1 P þS Ulus et al. (1998); Ilcol et al. (2002e)
After (6 h) childbirth 9.9–10.8 P þS Ulus et al. (1998); Ilcol et al. (2002e)
Before race 9.6–19.2 P Conlay et al. (1986, 1992); Buchman et al. (1999,
After (1–24 h) race 6.2–14.1 P Conlay et al. (1986, 1992); Buchman et al. (1999,
End‐stage renal disease
32.5–44.2 P þS Buchman et al. (2000b); Ilcol et al. (2002a, b)
Before surgery 10.7–12.3 P þS Ulus et al. (1998); Ilcol et al. (2002d, 2004, 2006)
After (1–24 h) surgery 5.6–8.8 P þS Ulus et al. (1998); Ilcol et al. (2002d, 2004, 2006)
Alzheimer’s disease patients 10.4–11.1 P Greenwald et al. (1985); Kanof et al. (1985)
Bipolar‐manic patients 12.1–12.3 P Jope et al. (1980); Stoll et al. (1991)
Alzheimer’s disease patients 40.7–48.1 RBC Greenwald et al. (1985); Kanof et al. (1985)
Bipolar‐manic patients 23.9–106 RBC Jope et al. (1980, 1984, 1985a, 1986); Stoll et al.
Unipolar‐depressed patients 17–51 RBC Wood et al. (1983); Jope et al. (1985b); Stoll et al.
Schizophrenic patients 16.7–90 RBC Kuchel et al. (1984); Jope et al. (1985b); Stoll et al.
Patients under lithium
277–626 RBC Jope et al. (1980, 1984, 1985a, 1986); Stoll et al.
Healthy adults 15.8–22.6 Blood Hasegawa et al. (1982); Danne et al. (2003)
Patients with noncardiac
19.9 0.7 Blood Danne et al. (2003)
Patients with unstable
24.5–47.4 Blood Danne et al. (2003)
Note: Values are given as the range of the means from the cited references, or as the mean SEM
P, plasma; S, serum; RBC, red blood cell; Blood, whole blood
Choline and its products acetylcholine and phosphatidylcholine 3.2 13
One molecule of choline oxidized through the respiratory chain yields two molecules of mitochondrial ATP
(Lin and Wu, 1986). Choline dehydrogenase has been cloned from rat liver mitochondria using a cDNA
formed from the enzyme’s terminal amino acid sequence (Huang and Lin, 2003). The enzyme is most active
in liver and kidney (Bernheim and Bernheim, 1933; Mann and Quastel, 1937), and, as discussed earlier, only
negligible choline dehydrogenase activity is observed in brain (Haubrich et al., 1979; Haubrich and Gerber,
1981). Recent estimates of choline dehydrogenase’s K
for choline—140–270 mM (Zhang et al., 1992)—
suggest values that are substantially lower than those estimated earlier (5–7 mM; Rendina and Singer, 1959;
Tsuge et al., 1980; Haubrich and Gerber, 1981). However, this revised K
is still high compared with actual
liver choline concentrations (60–230 mM; Sundler et al., 1972; Haubrich and Gerber, 1981), suggesting that
treatments that increase hepatic choline levels also increase its rate of degradation. Very high portal venous
choline concentrations (2.5 mM; Zeisel et al., 1980b) produced experimentally in studies on isolated
perfused livers can fully saturate the enzyme.
Betaine aldehyde dehydrogenase (EC 184.108.40.206) furtheroxidizes betaine aldehyde to betaine (>Figure 3.2‐2).
This enzyme, found both in cytoplasm and mitochondria (Wilken et al., 1970; Pietruszko and Chern,
2001), uses NAD as a cofactor (Wilken et al., 1970). Its K
for betaine aldehyde (118 mM in rat liver
mitochondria, Chern and Pietruszko, 1995; 123 mM in rat liver cytoplasm, Vaz et al., 2000; 182 mMin
human liver cytoplasm, Vaz et al., 2000) is probably substantially higher than actual in vivo concentrations
of the aldehyde; hence, the betaine aldehyde formed when hepatic choline levels rise is rapidly metabolized.
Choline’s oxidation occurs through two irreversible reactions; hence, betaine cannot be reduced back to
choline. Choline administration enhances betaine formation both in vivo (Haubrich et al., 1975; Zeisel and
Wurtman, 1981) and in vitro (Wilken et al., 1970; Zeisel et al., 1980b; Zhang et al., 1992). In isolated
perfused liver, betaine accounts for 60% of the labeled metabolites of [methyl‐
C]choline at concentra-
tions of 5–125 mM (Zeisel et al., 1980b). Betaine is used by liver as a source of methyl groups for the
generation of methionine from homocysteine, a process catalyzed by the enzyme betaine homocysteine
methyltransferase (EC 220.127.116.11; Klee et al., 1961). The importance of this pathway (and thus of choline as a
source of methyl groups) is probably increased when other methylating pathways have been compromised
by ethanol ingestion, drugs, or nutritional deﬁciencies affecting folic acid, pyridoxine, or vitamin B12
(Barak and Tuma, 1983). Thus, treatment with betaine could be beneﬁcial in early‐stage alcoholic liver
injury (Barak et al., 1996; Kharbanda et al., 2005).
Biosynthesis of sphingomyelin (SM) from phosphatidylcholine (PC) and ceramide
14 3.2 Choline and its products acetylcholine and phosphatidylcholine
2.3 Effects of Physiologic or Pathologic States on Plasma Choline
In adult humans, plasma (or serum) concentrations of free choline are maintained around 10 mM after
a 6–24 h fast (>Table 3.2‐4), and increased by up to 50 mM postprandially, depending on the choline
contents of the consumed food (Hirsch et al., 1978; Zeisel et al., 1980c). Basal plasma concentrations of free
choline decrease by about 25–30% in human subjects undergoing a 1‐week fast (Savendahl et al., 1997;
da Costa et al., 2004) or consuming a choline‐deﬁcient diet (<50 mg/day) for 3–6 weeks (Zeisel et al., 1991;
da Costa et al., 2006). Prolonged consumption of such diets can be associated with liver or muscle damage
(da Costa et al., 2004, 2006).
A number of physiologic or physiopathological circumstances (e.g., infancy, pregnancy, lactation,
surgery, parturition, marathon running, hemodialysis, and end‐stage renal disease) can alter baseline
plasma choline concentrations in humans (>Table 3.2‐4). These concentrations are much higher in
newborns (about 40 mM) than in adult humans (Zeisel et al., 1980a; Zeisel and Wurtman, 1981; Buchman
et al., 2001; Ilcol et al., 2002e, 2005a); a similar age relationship has been described in rats (Zeisel et al.,
1980a; Zeisel and Wurtman, 1981) and rabbits (Zeisel et al., 1980a). In rats, the high postnatal choline
concentrations fall to adult levels during the ﬁrst 3 weeks of postnatal life (Zeisel and Wurtman, 1981); in
humans, this occurs by 2 years of age (Ilcol et al., 2005a).
In humans, serum‐free choline levels rise gradually during pregnancy (Ilcol et al., 2002e; Velzing‐Aarts
et al., 2005) reaching 15–20 mM at term (Ulus et al., 1998; Ilcol et al., 2002e, 2005a); then they decrease by
35–40% during the 12–20 h after delivery (Ulus et al., 1998; Ilcol et al., 2002e). Like pregnancy, breast‐
feeding is associated with elevated serum‐free choline levels, in the mothers, rising to 15–20 mM during a
180‐day period of lactation. In both pregnancy and lactation, considerable amounts of free choline are
transferred from the mother to the fetus or breast‐fed infant, through the placenta or the breastmilk, and
used for growth and development (e.g., as a precursor of membrane phosphatides; Zeisel, 2006). The high‐
serum‐free choline concentrations observed in pregnant and lactating women may reﬂect a process for
promoting fetal and infant growth at the expense of depleting the mother’s choline stores (Zeisel et al.,
In patients with end‐stage renal disease, plasma‐free choline levels are several‐folds higher than those in
control subjects (Rennick et al., 1976; Buchman et al., 2000b; Ilcol et al., 2002a, b) or in patients who have
successfully undergone renal transplantation (Ilcol et al., 2002a). Considerable amounts of choline are lost
into hemodialysates (Rennick et al., 1976; Buchman et al., 2000b; Ilcol et al., 2002a) or peritoneal dialysates
(Ilcol et al., 2002b), but plasma‐free choline levels decrease only slightly during hemodialysis (Rennick et al.,
1976; Buchman et al., 2000b; Ilcol et al., 2002a).
Several studies have shown that plasma‐free choline concentrations decrease signiﬁcantly by about
25–40% after prolonged exercise, e.g., running a marathon (Conlay et al., 1986, 1992; Buchman et al., 1999,
2000a), and remain depressed for at least 48 h after the race (Buchman et al., 1999).
Serum‐free choline concentrations decrease by 20–45% during (Ulus et al., 1998; Ilcol et al., 2002d) and
after surgery, in humans (Ulus et al., 1998; Ilcol et al., 2002d, 2004, 2006) or dogs (Ilcol et al., 2003b). This
phenomenon is a response to surgical stress and is inversely correlated with the stress‐induced elevations in
serum cortisol, adrenocorticotropic hormone (ACTH), prolactin, and b‐endorphin (Ilcol et al., 2002a). The
magnitude and duration of surgery‐induced declines in serum choline depend on the severity and the type
of surgery (Ulus et al., 1998; Ilcol et al., 2002d, 2003b, 2005b). Thus, free choline concentrations return to
presurgical values within 24 or 48 h after a cesarean section (Ilcol et al., 2002e), or a transurethral
prostatectomy (Ulus et al., 1998), but require 72 h to do so after abdominal surgery (Ilcol et al., 2003b)
or 96 h after coronary artery bypass surgery (Ilcol et al., 2004, 2006) or removal of a brain tumor (Ilcol et al.,
2004). The decline in serum‐free choline concentration associated with surgery can be mimicked in dogs by
the administration of methylprednisolone (Ilcol et al., 2003b).
Perhaps paradoxically, plasma and whole blood choline concentrations reportedly increase signiﬁcantly
in patients with acute coronary syndromes (Danne et al., 2003, 2005).
Choline and its products acetylcholine and phosphatidylcholine 3.2 15
3 Choline in the Brain
Because choline is, by virtue of its quaternary nitrogen atom, relatively polar, it had generally been assumed
(Ansell and Spanner, 1971; Diamond, 1971) that plasma choline was unavailable to the brain. Moreover, as
brain cells were also thought to be incapable of synthesizing choline de novo, the ability of cholinergic
neurons to maintain the intracellular choline concentrations needed for ACh synthesis was usually
attributed either to an extraordinarily effective reuptake mechanism, described later, for reutilizing virtually
all the choline formed from the hydrolysis of ACh, or, less likely, to the uptake into brain of circulating PC
or lyso‐PC (Illingworth and Portman, 1972; Kuhar and Murrin, 1978). In addition, since the poor afﬁnity of
ChAT, the enzyme that catalyzes choline’s conversion to ACh for choline made it likely that intracellular
choline concentrations would control brain ACh synthesis; it was broadly conjectured that choline’s high‐
afﬁnity uptake from the synaptic cleft controlled ACh synthesis (cf., Taylor and Brown, 2006).
It is no longer held that the brain choline levels are sustained solely by the high‐afﬁnity uptake of free
choline from synapses, or that variations in this uptake are normally responsible for observed variations in
brain choline levels. Choline molecules (but not those of PC or lyso‐PC; Pardridge et al., 1979) do readily
cross the BBB (Cornford et al., 1978), and brain cells do indeed synthesize choline de novo (Blusztajn and
Wurtman, 1981). Physiological variations do occur in choline levels within brain neurons; however, these
result principally from changes in plasma choline concentrations after eating choline‐rich foods, or from
choline’s metabolism. It is possible in laboratory studies to make the reuptake of intrasynaptic choline
become the limiting factor controlling ACh synthesis, for example, by giving a drug‐like hemicholinium‐3
(HC3), which blocks the reuptake process. However, no food constituents or endogenous compounds have
ever been shown to share this ability. It is possible that the density of choline‐reuptake sites in nerve
terminals may be modulated by the rate of ACh release (Taylor and Brown, 2006); however, variations in the
rate of ACh release have not been demonstrated to affect the efﬁciency of choline reuptake.
Mammalian brains contain choline as the free base; as such water‐soluble phosphorylated metabolites
as phosphocholine and GPC (Nitsch et al., 1992), and as constituents of membrane phospholipids
including PC, SM, and lyso‐PC. Free choline levels in the brains of humans and rats reportedly vary
between 36–44 mM (Ross et al., 1997) and 30–60 mM (Stavinoha and Weintraub, 1974; Klein et al., 1993),
whereas PC and SM levels are orders of magnitude higher (2000–2500 mM and 250 mM, respectively;
Marshall et al., 1996). These high levels reﬂect the ubiquity of phospholipids, and the numerous essential
roles they mediate when they form membranes. Membrane phospholipids also serve as reservoirs for
choline and for such ‘‘second messenger’’ molecules as DAG, AA, inositol trisphosphate (IP3), and
3.1 Sources of Brain Choline
Free choline molecules in the brain derive from four known sources such as uptake from the plasma;
liberation from the PC in brain membranes; high‐afﬁnity uptake from the synaptic cleft after ACh released
from a cholinergic terminal has been hydrolyzed; and, probably to a minor extent, the breakdown of newly
synthesized PC formed from the methylation of PE.
3.1.1 Uptake of Circulating Choline into the Brain
The brain can obtain circulating choline and various other circulating nutrients (e.g., neutral and basic
amino acids, glucose, adenine, or adenosine; Pardridge and Oldendorf, 1977; Pardridge, 1986) via two
routes: Small amounts can pass from the blood to the cerebrospinal ﬂuid through the action of a speciﬁc
transport protein, organic cation transporter 2 (OCT2), which is present in cells that line the choroid plexus
(CP) (Sweet et al., 2001). However, orders of magnitude more pass bidirectionally between the blood and
16 3.2 Choline and its products acetylcholine and phosphatidylcholine
the brain’s extracellular ﬂuid (ECF) by facilitated diffusion. This process is catalyzed by a different transport
protein, not yet cloned, which is localized within the endothelial cells that line the brain’s capillaries
(Oldendorf and Braun, 1976; Pardridge and Oldendorf, 1977; Cornford et al., 1978). Its action is indepen-
dent of sodium, and can be blocked by HC3.
Studies using an in situ brain perfusion technique, or a cell line of immortalized endothelial micro-
vessels from rat brain (RBE4), have demonstrated the existence of a transport protein with a relatively low
for choline [(39–42 mM; Allen and Smith, 2001) or (20 mM; Friedrich et al., 2001)] which could mediate
choline’s bidirectional ﬂux across the BBB. Other investigators using other experimental systems had
proposed substantially higher K
s for endothelial choline transport, i.e., 220–450 mM (Oldendorf and
Braun, 1976; Pardridge and Oldendorf, 1977; Cornford et al., 1978; Mooradian, 1988). The differences
among the afﬁnities noted in these studies might, as discussed later, reﬂect the different methods used for
their measurement. However, in any case, the protein would still be unsaturated at physiological plasma
choline concentrations, and its net activity is still affected by variations in these concentrations. It might
constitute a kind of pore through which choline can pass in either direction, based on the gradient between
its blood and brain levels (Klein et al., 1990). Hence when plasma choline levels have been elevated, for
example by eating a choline‐rich meal (e.g., to 50 mM in the rat; Zeisel et al., 1980a), choline tends to enter
the brain, but when plasma choline levels are low its ﬂux is in the opposite direction. It has been estimated
that the plasma choline concentration in rats required in order for the net choline ﬂux to be from blood to
brain is about 15 mM; below this concentration, net choline ﬂux presumably is from brain to blood (Klein
et al., 1990).
No endogenous circulating compound has been shown to compete effectively with choline for
facilitated diffusion across the BBB. Very high concentrations of carnitine and spermidine, compared
with those in the blood, can reduce brain uptake of choline by 20–25% (Cornford et al., 1978). One
drug, diethylaminoethanol, apparently does block BBB choline uptake, and has been used to lower brain
choline levels and thereby suppress ACh synthesis (Cornford et al., 1978; Millington et al., 1978). Lithium
ion, given acutely (Cornford et al., 1978) or chronically (Millington et al., 1978), may also block BBB
choline uptake. However, lithium also blocks choline’s efﬂux from brain to blood, thus producing a net
increase in brain choline levels (Millington et al., 1978).
Once circulating choline has entered the brain’s ECF, it can be taken up into all cells by a low‐afﬁnity
transport protein (K
¼30–100 mM), or into cholinergic nerve terminals by a high‐afﬁnity uptake protein
¼0.1–10 mM) (Haga and Noda, 1973; Yamamura and Snyder, 1973; Blusztajn and Wurtman, 1983).
Both of these are described later. The high‐afﬁnity process, unlike the passage of choline across the BBB, is
3.1.2 Liberation from Membrane PC
The choline in membrane PC can be liberated through the actions of the phospholipase enzymes, described
earlier, which catalyze the hydrolysis of various bonds between PC’s three oxygen molecules and fatty acids
or its phosphate moiety (>Figure 3.2‐5). The activation of each of these enzymes is tightly regulated and, in
general, initiated by the interaction of a neurotransmitter or other biologic signal with a receptor coupled to
aG‐protein. For example, both the PLC enzymes (which act on PC to yield DAG and phosphocholine, or on
PI) and PLD (which acts on PC to yield phosphatidic acid and choline) are activated when ACh attaches to
M1 or M3 muscarinic receptors (Sandmann and Wurtman, 1990, 1991; Sandmann et al., 1991). The DAG
generated by PLC activates a family of protein kinase (PK) enzymes that phosphorylate various proteins,
including those that control the metabolism of the amyloid precursor protein (APP) to form either soluble
APP or the A‐beta peptides (Hung et al., 1993; Nitsch et al., 1994; Slack et al., 1997; Slack and Wurtman,
The release of choline from PC can also be enhanced, and its reincorporation into PC is diminished by
sustained neuronal depolarization (Farber et al., 1996). This process has been termed ‘‘autocannibalism’’
when some of the choline is diverted for the synthesis of ACh (Blusztajn et al., 1986; Ulus et al., 1989).
Choline and its products acetylcholine and phosphatidylcholine 3.2 17
Autocannibalism may, by decreasing the quantities of phosphatide molecules, and thus of neuronal
membranes, underlie the particular vulnerability of cholinergic neurons in certain diseases (Blusztajn
et al., 1986; Ulus et al., 1989). It is not known whether the accelerated breakdown of PC associated with
sustained neuronal depolarization results from changes in ion ﬂux or requires the release of local neuro-
transmitters and activation of particular receptors. The depletion of membrane PC and other phospha-
tides—including those not containing choline—caused by frequent or sustained depolarizations can be
diminished or blocked entirely, and the release of ACh is enhanced by providing the brain with supplemen-
tal choline (Ulus et al., 1989).
3.1.3 Reutilization of Choline Formed from Hydrolysis of Acetylcholine
ACh released into synapses is rapidly hydrolyzed to free choline and acetate. This process terminates the
neurotransmitter’s physiologic actions, i.e., its ability to combine with and activate its pre‐or postsynaptic
muscarinic or nicotinic receptors. (The inactivation of ACh differs from that of other aminergic transmit-
ters, e.g., dopamine and serotonin, in which it involves a chemical change in the neurotransmitter molecule,
and not simply physical removal of that molecule from the synaptic cleft by reuptake into its nerve terminal
of origin.) The enzymes that catalyze ACh hydrolysis, the acetylcholinesterases (EC 18.104.22.168; AChE), are
particularly abundant within the cholinergic synapse; they are synthesized in the cholinergic neuron and
secreted into the synapse, along with ACh, when the neuron is depolarized. A related enzyme, butyrylcho-
linesterase (EC 22.214.171.124; BuChE), synthesized in the liver and present in plasma, probably functions to
inactivate potentially toxic dietary esters but not intrasynaptic ACh: It is active in the nervous system during
development, but not thereafter, and mutant animals lacking the BuChE gene—in contrast to those lacking
AChE—apparently fail to exhibit neurologic symptoms (Taylor and Radic, 1994; Giacobini, 2003).
Most of the free choline liberated by the intrasynaptic hydrolysis of ACh is taken back up into its nerve
terminal of origin by the high‐afﬁnity choline transporter (CHT) described later, and either reacetylated to
form ACh or phosphorylated for ultimate conversion to membrane PC (Ulus et al., 1989).
3.1.4 De Novo Synthesis of Phosphatidylcholine and Choline
As described earlier, brain cells—including nerve terminals (Holbrook and Wurtman, 1988)—contain all
the enzymes needed to synthesize PC from ethanolamine (>Figure 3.2‐3) or from PS. These include the
Kennedy cycle enzymes that convert ethanolamine to PE (Spanner and Ansell, 1979), PS decarboxylase,
which forms PE from PS (Butler and Morell, 1983), and the enzymes (PEMT1 and PEMT2), which
methylate PE (Crews et al., 1980).
4 Brain Proteins that Interact with Choline
Free choline is known to interact with two brain enzymes and four transport proteins, as well as various
receptors for ACh.
The two enzymes are ChAT and CK—which catalyze, respectively, the transformations of choline to
ACh within cholinergic terminals, and to phosphocholine within all cells.
The four transport proteins include two that move choline across the BBB, i.e., the facilitated‐diffusion
site in brain capillaries through which choline passes, bidirectionally, between the plasma and the brain’s
ECF, and the organic cation transporter that carries plasma choline across the CP and into the cerebrospinal
ﬂuid; and two that enable the choline in brain ECF to enter cells, i.e., the low‐afﬁnity uptake site that
catalyzes choline’s uptake into all brain cells, and the high‐afﬁnity uptake site that transports intrasynaptic
choline into the presynaptic terminals of cholinergic neurons.
18 3.2 Choline and its products acetylcholine and phosphatidylcholine
The cholinergic receptors, which choline can also activate, include both nicotinic and muscarinic
varieties. This section describes the properties of each of these proteins, and the consequences of their
interactions with choline.
4.1.1 Choline Acetyltransferase
ChAT (acetyl‐CoA: choline‐O‐acetyltransferase, EC 126.96.36.199) mediates a single reaction, the transfer of an
acetyl group from acetyl‐coenzyme A (acetyl‐coA) to choline, which thereby generates the neurotransmitter
ACh in cholinergic neurons.
ChAT, a single‐stranded globular protein, is encoded by a single gene with, in humans, six distinct
transcripts formed from the alternative splicing of ﬁve noncoding exons (Misawa et al., 1992, 1997; Oda
et al., 1992; Robert and Quirin‐Stricker, 2001) Polymorphism among these transcripts is apparently limited
to their 50‐untranslated regions. In humans, four of the six transcripts (designated as H, R, N1, and N2)
translate to the same 69‐kD protein (Misawa et al., 1992, 1997; Oda et al., 1992; Robert and Quirin‐Stricker,
2001). The ﬁfth and sixth transcripts, designated as M and S, have two translation sites and yield, besides the
69‐kD enzyme, 82‐kD and 74‐kD forms of ChAT, respectively. The 82‐kD ChAT differs from the 69‐kD
form in that it has an aminoterminal extension with 118 amino acid residue (Oda et al., 1992; Misawa et al.,
1997). Physiological roles for the 74‐kD and 82‐kD forms of ChATremain to be elucidated, and indeed it is
not clear that these larger forms of human ChAT actually are synthesized in vivo (Oda, 1999).
ChAT probably exists in at least two forms within cholinergic nerve terminals—a soluble form (80–90%
of the total enzyme activity) and a membrane‐associated form (10–20%; Benishin and Carroll, 1981; Salem
et al., 1994; Pahud et al., 1998). These two forms exhibit different physicochemical and biochemical
properties (Benishin and Carroll, 1983; Eder‐Colli et al., 1986; Pahud et al., 2003). The soluble form is
hydrophilic, and the membrane‐bound form is amphiphilic (Benishin and Carroll, 1983; Eder‐Colli et al.,
1986; Pahud et al., 2003). Soluble ChAT has higher afﬁnities for both of its substrates, choline and acetyl‐
CoA, when assayed at low ionic strength (K
for choline 350 mM; for acetyl‐CoA 2.5 mM) than that when
assayed at higher ionic strengths (K
for choline 6700 mM; for acetyl‐CoA 77 mM; Rossier, 1977). The
activity of ChAT in crude synaptosomal preparations (presumably representing a mixture of the soluble and
membrane‐bound forms) also varies with ionic strength; the afﬁnity of synaptosomal ChAT for choline
appears to be greater than that of soluble ChAT (K
¼22 mM at low ionic strength and 540 mM at high ionic
strength; Rossier, 1977). In any case, ChAT is invariably unsaturated with choline at the choline concentra-
tions that could exist within nerve terminals (Tucek, 1990), indicating that ChAT is in kinetic excess (Hersh,
1982; Tucek, 1990), and that its substrate‐saturation, not its levels, is rate limiting in ACh synthesis. Both
choline and acetyl‐CoA (Rossier, 1977; Hersh, 1982; Tucek, 1990) levels can affect the rate at which ACh is
There is also evidence that phosphorylation and dephosphorylation of ChAT can alter its catalytic
activity, subcellular distribution, and interactions with other cellular proteins (see review of Dobransky and
Rylett, 2005). ChAT is a substrate for multiple PKs; 69 kDa ChAT is phosphorylated by PK‐C, PK‐CK2, and
/calmodulin‐dependent PK‐II (CaM‐kinase) but not by PK‐A, whereas 82 kDa ChAT is phosphory-
lated by PK‐C and CaM‐kinase (Dobransky et al., 2000, 2001). ChAT is differentially phosphorylated by
PK‐C isoforms on four of its serine residues (Ser‐440, Ser‐346, Ser‐347, and Ser‐476) and one threonine
residue (Thr‐255); this phosphorylation is hierarchical, such that phosphorylation at Ser‐476 is required in
order for the other serines to become phosphorylated (Dobransky et al., 2004). Phosphorylation at some
but not all of those sites (Ser‐476 with Ser‐440 and Ser‐346/347; Dobransky et al., 2004) affects basal ChAT
activity. Phosphorylation of ChAT by PK‐C alone can double the enzyme’s activity, whereas coordinated
phosphorylation of ChAT at threonine 456 (by CaM‐kinase II) and serine 440 (by PK‐C) can treble ChAT
activity (Dobransky et al., 2003). Whether the phosphorylation and dephosphor ylation of ChAT also alter
the enzyme’s afﬁnities for choline or acetyl‐CoA in intact cells is clear.
Choline and its products acetylcholine and phosphatidylcholine 3.2 19
4.1.2 Choline Kinase
CK (ATP:choline phosphotransferase; EC 188.8.131.52) catalyzes the ﬁrst phosphorylation reaction in the
Kennedy cycle of PC synthesis (>Figure 3.2‐4); ATP is the phosphate donor and the presence of Mg
required (Wittenberg and Kornberg, 1953). CK can also catalyze the phosphorylation of ethanolamine, as
well as N‐monomethylethanolamine and N,N‐dimethylethanolamine (Ishidate et al., 1985; Porter and Kent,
1990; Uchida and Yamashita, 1990); however, a separate ethanolamine kinase enzyme exists, demonstrated
by cloning cDNA from human liver (Lykidis et al., 2001). CK is mainly cytosolic but is also associated with
particulate (membrane‐bound) fractions of rat striatum (Reinhardt and Wecker, 1983). The enzyme has
been puriﬁed to homogeneity from various rat tissues (Ishidate et al., 1985; Porter and Kent, 1990),
including brain (Uchida and Yamashita, 1990). HC3 (Ansell and Spanner, 1974) and ADP (Burt and
Brody, 1975) inhibit CK activity in vitro, whereas high concentrations of the polyamines spermine and
spermidine (Uchida and Yamashita, 1990) enhance its activity. Several isoforms of CK exist in brain and
other tissues, differentiable by their subunit masses (Porter and Kent, 1990; Uchida and Yamashita, 1990)
and by cloning and expression studies (Uchida and Yamashita, 1992; Uchida, 1994; Aoyama et al., 1998). At
least as three isoforms (CK‐a1, CK‐a2, and CK‐b), encoded by two separate genes termed ck‐aand ck‐b
(Aoyama et al., 1998, 2000, 2004), are now recognized. The latter resides on chromosome 22q13 in humans
(Froguel and McGarry, 1997); the locus of the former awaits determination. These isoforms may not be
active as monomers, but become active on forming dimeric or oligomeric structures (Aoyama et al., 2004).
In some circumstances, CK activity may be rate limiting in PC synthesis; for example, a 3.5‐fold increase
in CK activity in livers of rats deﬁcient in essential fatty acids was accompanied by a parallel increase in PC
synthesis (Infante and Kinsella, 1978). Similar relationships have been described in livers of estrogen‐treated
roosters (Vigo and Vance, 1981) or quiescent murine 3T3 cells in culture (Warden and Friedkin, 1985).
However, it is probably not the activity of CK per se, but rather its degree of substrate saturation that affects
the rate of PC synthesis. The K
of CK for choline in rat brain is 14–134 mM (Uchida and Yamashita, 1990;
Cao and Kanfer, 1995); this value is 32–310 mM in rabbit brain (Haubrich, 1973). An even higher K
(2.6 mM) was described by Spanner and Ansell (1979) who assayed the enzyme at a more physiological pH
(7.5) than that customarily used (pH ¼9.0); this allowed phosphocholine, CK’s reaction product, to be
assayed without ﬁrst being hydrolyzed. Hence, CK is unsaturated with choline at normal brain choline
concentrations (30–60 mM), and the production of phosphocholine through CK, like that of ACh by ChAT,
is controlled principally by brain choline levels (Millington and Wurtman, 1982; Cohen et al., 1995).
4.2 Transport Proteins
4.2.1 Facilitated‐Diffusion Carrier at Blood–Brain Barrier
A transport protein that mediates the bidirectional facilitated‐diffusion of choline at the BBB has been
identiﬁed (Oldendorf and Braun, 1976; Pardridge and Oldendorf, 1977; Cornford et al., 1978). This protein
does not require metabolic energy or sodium ﬂux and cannot maintain a concentration gradient.
The CHT at the BBB, as discussed earlier, might allow bidirectional passage of choline based on the
gradient between its blood and brain concentrations (Klein et al., 1990). A blood choline concentration of
15 mM has been estimated to be required for choline inﬂux to the rat’s brain to predominate; below this
concentration, choline efﬂux predominates (Klein et al., 1990).
As discussed earlier, the BBB CHT’s K
for choline in vivo is estimated as 220–450 mM (Oldendorf and
Braun, 1976; Pardridge and Oldendorf, 1977; Cornford et al., 1978; Mooradian, 1988). Hence, it is
unsaturated at physiologic plasma choline concentrations (10 mM). The K
of BBB CHT for choline
has been demonstrated as 39–42 mM in a recent study using an in situ brain perfusion technique (Allen and
Smith, 2001); the higher afﬁnity of the BBB transporter for choline found in this study, compared with
previous data, probably reﬂects methodological differences. In vitro assays, using cell lines of immortalized
rat (RBE4; Friedrich et al., 2001) and mouse (MBE4; Sawada et al., 1999) brain endothelial microvessels,
have demonstrated the existence of a transport protein with a relatively low K
for choline (20 mM in both
20 3.2 Choline and its products acetylcholine and phosphatidylcholine
studies). However, these studies investigated uptake only at the luminal side of the endothelial cells, and not
at both luminal and abluminal sides as was investigated in previous in vivo studies. The CHT at the BBB has
not yet been cloned and awaits further characterization. A perhaps‐related transporter for carnitine, the
organic cation/carnitine transporter OCTN2 (Tamai et al., 1998), has been cloned from RBE4 cells (Friedrich
et al., 2003). Carnitine uptake through this transporter is not blocked by choline (Tamai et al., 1998).
4.2.2 Choroid Plexus Choline Transporter
Much less blood choline is transported through the CP epithelium to the cerebrospinal ﬂuid than through
the BBB carrier to the brain’s ECF, because the surface area of the CP epithelium is much smaller than that
of the BBB epithelium (Pardridge, 2001). Ventricular choline transport, in the rat, is mediated by one of the
OCT proteins, OCT2 (Sweet et al., 2001). The three subtypes of OCTs (OCT1–3) have been isolated from
rat (Grundemann et al., 1994; Okuda et al., 1996; Kekuda et al., 1998), mouse (Schweifer and Barlow, 1996;
Mooslehner and Allen, 1999), and human tissues (Gorboulev et al., 1997; Zhang et al., 1997; Grundemann
et al., 1998). OCTs are transmembrane proteins with 12 membrane‐spanning domains (Koepsell et al.,
2003). Transport of a cation by an OCT protein is electrogenic, Na
‐independent, and reversible with
respect to direction (Koepsell and Endou, 2004). In the human, genes encoding OCT1–3 have been found
on chromosome 6 (6q26–6q27; Koehler et al., 1997, 2003). The hOCT2 protein is expressed in kidney
(Gorboulev et al., 1997) and brain (Busch et al., 1998); however, its localization in human brain ventricles,
and its possible transport activity remain to be established.
4.2.3 High‐Afﬁnity Uptake Protein in Cholinergic Terminals
A saturable, sodium‐and energy‐dependent, HC3‐sensitive, high‐afﬁnity CHT has been demonstrated in
synaptosomes (Yamamura and Snyder, 1972, 1973; Guyenet et al., 1973; Haga and Noda, 1973). The K
this transporter for choline is 0.1–10 mM (Guyenet et al., 1973; Haga and Noda, 1973; Yamamura and
Snyder, 1973; Blusztajn and Wurtman, 1983). Choline uptake through the CHT is competitively inhibited
by nanomolar concentrations of HC3 (K
: 10–100 nM; Yamamura and Snyder, 1972; Haga and Noda, 1973;
Kuhar and Murrin, 1978).
The high‐afﬁnity CHT protein, made up of two polypeptides with molecular masses of 58 and 35 kDa,
has been identiﬁed and partially puriﬁed from rat corpus striatum (Rylett et al., 1996). cDNAs from rat
(rCHT1; Okuda et al., 2000), mouse (mCHT1; Apparsundaram et al., 2001), and human (hCHT1;
Apparsundaram et al., 2000) have also been isolated, cloned and expressed. These studies have shown
that CHT1 does not belong to the neurotransmitter transporter family, but rather to the sodium‐dependent
glucose transporter family (SLC5, in which CHT1 is designated as SLC5A7) (Apparsundaram et al., 2000;
Okuda et al., 2000; Okuda and Haga, 2003). CHT1 protein has 13 transmembrane domains (Apparsun-
daram et al., 2000; Okuda et al., 2000). The human CHT1 gene is localized on chromosome 2q12
(Apparsundaram et al., 2000).
High‐afﬁnity choline transport occurs predominantly into terminals of cholinergic neurons (Misawa
et al., 2001). Using antibodies raised against CHT1, CHT‐immunoreactive cells have been shown to be
widely distributed throughout the rat, primate, and human central nervous systems (Misawa et al., 2001;
Kus et al., 2003). Primate cerebellums contain numerous CHT‐immunoreactive cells (Kus et al., 2003), and
mouse cerebellum expresses CHT1 mRNA, particularly during development (Berse et al., 2005). CHT1 is
also present in terminals and in those of peripheral motor neurons (Lips et al., 2002; Nakata et al., 2004)
and of parasympathetic neurons to the tongue (Lips et al., 2002). CHT1 is not expressed in glial cells (Inazu
et al., 2005) but, contrary to what had been believed, is expressed in such nonneuronal cells as rat trachea
(Pfeil et al., 2003), rat and human arteries (Lips et al., 2003), and skin (Haberberger et al., 2002). The
function of CHT1 in these tissues awaits determination.
Within neurons, CHT immunoreactivity is detectable in cell soma, proximal dendrites, axons and,
particularly, axon terminals (Kus et al., 2003). Within the terminals, the CHT1 protein is especially
Choline and its products acetylcholine and phosphatidylcholine 3.2 21
abundant in plasma membrane, synaptic vesicles, and endosomal vesicles (Ferguson et al., 2003; Ribeiro
et al., 2003; Ferguson and Blakely, 2004). In motor neurons of diaphragm, the CHT is mainly (>90%)
concentrated within synaptic vesicles, rather than in the presynaptic membrane itself (Nakata et al., 2004).
These vesicles may store CHT1 in the resting state, and the protein may migrate to the synaptic membrane
during depolarization (Ribeiro et al., 2003, 2005).
Activity‐dependent modulation of CHT1 has been described in studies using several different experi-
mental systems, for example electrical or pharmacological stimulation of cholinergic neurons in vitro
(reviewed in Ferguson and Blakely, 2004). The capacity and density of CHTs are apparently increased
in medial prefrontal cortices of rats performing attentional tasks (Apparsundaram et al., 2005). Neuronal
activity per se (Ferguson et al., 2003) might, by altering the phosphorylation state of CHT1 protein (Gates
et al., 2004), enhance the transfer of CHT1 into and out of vesicles, thus modulating its activity. NGF,
which can upregulate CHT1 through a PI3K‐dependent process, might similarly inﬂuence its activity (Berse
et al., 2005).
4.2.4 Low‐Afﬁnity Cellular Uptake Protein
A nonsaturable, Na
‐independent, high‐capacity, and low‐afﬁnity CHT has also been identiﬁed (Haga and
Noda, 1973; Yamamura and Snyder, 1973). Not surprisingly—since all cells need choline for phospholipid
synthesis—it appears to be ubiquitous in mammals and is found, in, for example, kidney (Bevan and Kinne,
1990), liver (Zeisel et al., 1980), and placenta (Grassl, 1994) as well as in brain synaptosomes (Ferguson
et al., 1991). The K
of the low‐afﬁnity transport protein for choline varies between 30 and 100 mM
(Dowdall and Simon, 1973; Haga and Noda, 1973; Yamamura and Snyder, 1973); it also is inhibited by HC3
s of about 40–50 mM (Haga and Noda, 1973) in brain and 100 mM in human placenta (Grassl, 1994).
Low‐afﬁnity choline transport has been suggested to be a carrier‐mediated process (Ferguson et al., 1991;
Inazu et al., 2005). As discussed later, CTL1, a member of the family of choline‐transporter‐like proteins
(Traiffort et al., 2005), has been proposed as mediating the low‐afﬁnity transport of choline into rat cortical
astrocytes (Inazu et al., 2005) and mouse cortical neurons (Fujita et al., 2006).
The choline‐transporter‐like proteins CTL1–CTL5 are encoded by ﬁve different genes, also labeled from
CTL1 to CTL5 (Traiffort et al., 2005). CTL1, a transmembrane protein with 10 transmembrane domains,
has been cloned and characterized from rat (rCTL1; O’Regan et al., 2000), human (hCTL1; Wille et al.,
2001), and mouse (mCTL1; Yuan et al., 2004) tissues. The human gene is located on chromosome 9q31.2
(Wille et al., 2001), and its protein product is expressed as two polypeptides, of 50 and 23 kDa, which have
been found in such tissues as brain, heart, small intestine, kidney, liver, lung, skeletal muscle, pancreas,
spleen, ovary, and testis (Yuan et al., 2006).
Another choline‐transporting system, the OCT proteins (members of the solute carrier family SLC22;
Koepsell et al., 2003; Koepsell and Endou, 2004), have also been implicated in low‐afﬁnity choline uptake.
For example, rat OCT1 (rOCT1), cloned from renal proximal tubule epithelial cells or hepatocytes and
expressed in Xenopus oocytes, can mediate low‐afﬁnity choline uptake (K
¼1.1 mM; Busch et al., 1996),
and human OCT1 (hOCT1) and human OCT2 (hOCT2) can mediate, respectively, hepatic and renal
choline transport (K
¼210 mM; Gorboulev et al., 1997).
It has not yet been determined whether the low‐afﬁnity CHT is one of the OCT proteins; CTL1
(perhaps more likely in brain; Inazu et al., 2005; Fujita et al., 2006); or even a different protein.
Choline in sufﬁciently high concentrations can directly activate both muscarinic (mAChRs) and the
nicotinic (nAChRs) acetylcholine receptors. The ﬁve muscarinic receptors (M1–M5) mediate slow meta-
bolic responses to ACh, and the nicotinic receptors, which are ligand‐gated ion channels, implement fast,
ACh‐mediated synaptic transmission in the CNS, ganglia, and neuromuscular synapses. The M1, M3, and
22 3.2 Choline and its products acetylcholine and phosphatidylcholine
M5 muscarinic receptors activate phospholipase C, thereby generating the second messengers IP3 and DAG
(Caulﬁeld and Birdsall, 1998); the M2 and M4 muscarinic receptors inhibit adenylate cyclase activity, thus
reducing intracellular cAMP levels, or can enhance the ﬂux of potassium and other ions through nonselec-
tive ion channels. The nicotinic receptors, pentameric structures made up of combinations of 17 known
individual subunits, increase the ﬂux of sodium into postsynaptic cells, thus increasing the likelihood of the
cells’ depolarization. Free choline concentrations in synaptic ﬂuid following neuronal depolarization
apparently have not been measured, and may or may not attain levels sufﬁcient to activate cholinergic
receptors under physiological circumstances. Much higher concentrations, produced experimentally, are
readily shown to activate the receptors in vitro.
Many years ago, it was noted that choline can produce ‘‘muscarine‐like’’ or ‘‘nicotine‐like’’ effects in
various peripheral tissues (Dale, 1914; Chang and Gaddum, 1933), voluntary muscles (Bacq and Brown,
1937; Hutter, 1952; Del Castillo and Katz, 1957), and autonomic ganglia (Feldberg and Vartiainen, 1934;
Kosterlitz et al., 1968; Krstic, 1972), with 1/20,000 to 1/714th the potency of ACh (Chang and Gaddum,
1933). Functional (Pomeroy and Raper, 1972; Ulus et al., 1979, 1988a; Holz and Senter, 1981) and receptor
binding studies (Speth and Yamamura, 1979; Palacios and Kuhar, 1979; Costa and Murphy, 1984; Ulus
et al., 1988), in which the choline presumably had not ﬁrst been acetylated to authentic ACh, identiﬁed the
effective choline concentrations needed to bind to AChRs and/or to produce ACh‐like biological responses.
Choline, acting as a direct muscarinic agonist, excited cortical neurons (Krnjevic and Reinhardt, 1979);
contracted isolated smooth muscle in rat stomach fundus (EC50¼0.41 mM), rat trachea (EC50¼1.7 mM),
rat urinary bladder (EC50¼10.9 mM) (Ulus et al., 1988), and guinea pig ileum (EC¼0.6 mM; Pomeroy
and Raper, 1972 or EC50¼0.20 mM; Ulus et al., 1979); and reduced the frequency at which isolated rat or
guinea pig right atrium beat spontaneously (Ulus et al., 1979, 1988). It also inhibited ACh release from
myenteric plexus‐longitudinal muscle preparations of guinea pig ileum (EC50¼0.3 mM) (Kilbinger and
Kruel, 1981), and inhibited [
H]‐quinuclidinyl benzilate binding to rat brain membranes (Placios and
Kuhar, 1979; Speth and Yamamura, 1979; Costa and Murphy, 1984; Ulus et al., 1988) and rat peripheral
tissues (Ulus et al., 1988). Choline’s potency for inhibiting [
H]‐quinuclidinyl benzilate binding was found
to vary among brain regions (K
¼0.46–3.5 mM) (Palacios and Kuhar, 1979; Speth and Yamamura, 1979;
Costa and Murphy, 1984; Ulus et al., 1988) and also in peripheral tissues (K
¼0.28–1.17 mM) (Ulus et al.,
1988). The wide range of variations in the muscarinic potency of choline and in its relative tissue selectivity
(Ulus et al., 1988) may result from its varying afﬁnities for mAChRs subtypes (M1–M5). Choline acts as a
full agonist on human mutant M1 receptors to stimulate phosphoinositide hydrolysis (EC50¼0.2 mM;
Huang et al., 1998), and on cloned human M1 receptor to stimulate nitric oxide synthesis and elevate
, at 0.1–1.0 mM concentrations (Carriere and El‐Fakanay, 2000).
By activating nicotinic receptors as a full agonist (Ulus et al., 1988) and/or a partial agonist (Holz and
Senter, 1981), choline stimulates catecholamine secretion from the vascularly perfused adrenal gland
(EC50¼2.1 mM; Ulus et al., 1988), and from primary cultures of bovine adrenal chromafﬁn cells (at
1–10 mM; Holz and Senter, 1981). It also competes with L‐[
H]‐nicotine for binding to membrane
preparations of rat brain (Costa and Murphy, 1984; Ulus et al., 1988) and peripheral tissues (Ulus et al.,
1988). The potency of choline in displacing L‐[
H]‐nicotine from brain nicotinic receptors varies within a
threefold range (K
¼379–1167 mM), and within a 1.5‐fold range for peripheral tissues (K
Ulus et al., 1988). Patch‐clamp studies have shown that choline interacts with nAChRsin a ‘‘subtype selective’’
and concentration‐dependent manner. At concentrations of 0.1–10 mM, choline acts as a full agonist on a7
nAChR (Mandelzys et al., 1995; Papke et al., 1996, 2000; Alkondon et al., 1997, 2000; Albuquerque et al., 1998;
Cuevas et al., 2000; Papke and Papke, 2002; Alkondon and Albuquerque,2006; Gonzales‐Rubio et al., 2006) or a
partial agonist on a3b4 nAChRs (Mandelzys et al., 1995; Papke et al., 1996; Albuquerque et al., 1998), a3b4*
nAChRs (Seddik et al., 2003), and a4b4 nAChRs (Zwart and Vijverberg, 2000). It desensitizes a7nAChRs at
10–100 mM concentrations (Mandelzys et al., 1995; Papke et al., 1996, 2000, 2002; Albuquerque et al., 1997;
Alkondon et al., 1997), inhibits a3b4* and a4b2* nAChRs at 10–1000 mM concentrations (Alkondon
and Albuquerque, 2006), and potentiates or inhibits a4b4nAChR‐mediated ACh currents at 10–300 mMor
1–30 mM concentrations, respectively (Zwart and Vijverberg, 2000).
Choline and its products acetylcholine and phosphatidylcholine 3.2 23
5 Utilization of Choline in Brain
All cells use choline to produce the PC and SM in their membranes. Cholinergic neurons also use choline
for an additional purpose, synthesis of their neurotransmitter, ACh. Both the PC and the ACh are ultimately
broken down to regenerate free choline, thus both of these compounds can also be considered ‘‘reservoirs’’
for free choline. The synthesis of PC (>Figure 3.2‐4) is initiated by the phosphorylation of choline,
catalyzed by an enzyme, CK, which forms phosphocholine by transferring a monophosphate group from
ATP to the hydroxyl oxygen of the choline. As described later, this phosphocholine then combines with
cytidine‐50‐triphosphate (CTP) to form cytidine‐50‐diphosphocholine (CDP‐choline), which, in turn,
combines with DAG to yield PC. The synthesis of ACh, catalyzed by the enzyme ChAT, involves a single
reaction, the transfer of an acetyl group from acetyl‐CoA, also to the hydroxyl oxygen of the choline. The
ACh is then stored, largely within synaptic vesicles, for future release.
Both CK and ChAT have low afﬁnities for their choline substrate: Their K
s in brain, which describe the
choline concentrations at which the enzymes operate at only half‐maximal velocity, may be as high as
2.6 mM (Spanner and Ansell, 1979) and 540 mM (Rossier, 1977), respectively, whereas brain choline levels,
as noted earlier, are only about 30–60 mM, and thus well below the concentrations that would probably be
needed to enable either enzyme to operate at maximal velocity. Hence, both of the enzymes are highly
responsive to treatments that raise or lower brain choline levels.
The ability of choline administration to increase the syntheses and brain levels of phosphocholine and
ACh was ﬁrst noted in 1982 (Millington and Wurtman, 1982) and 1975 (Cohen and Wurtman, 1975;
Haubrich et al., 1975). It had previously been shown that the synthesis and levels of another brain
neurotransmitter, serotonin, were increased if animals were given physiologic doses of its circulating
precursor, tryptophan (Fernstrom and Wurtman, 1971; Cansev and Wurtman, 2006). This was because
tryptophan hydroxylase, the enzyme that determines the overall rate at which tryptophan is converted to
serotonin, has a low afﬁnity for this substrate. Moreover, since ChAT’s afﬁnity for choline had also been
shown, in vitro, to be low, it seemed reasonableto enquire into whether choline’s ChAT‐mediated conversion
to ACh also was precursor‐dependent. Once this relationship was afﬁrmed, experiments soon followed
demonstrating the precursor‐dependence of phosphocholine synthesis (Millington and Wurtman, 1982).
Even though brain choline concentrations shared with those of tryptophan the ability to control the
rates at which the precursor is of used for neurotransmitter synthesis, the two precursors differed in an
important respect: Although tryptophan and choline are both used by certain neurons for two purposes:
tryptophan for conversion to serotonin and incorporation into proteins, and choline for conversion to ACh
and incorporation into phospholipids, in the case of tryptophan these two processes are segregated into
different parts of the neuron—the nerve terminal and perikaryon, respectively—whereas for choline both
can take place within the nerve terminal, inasmuch as that structure contains both ChAT and CK. Hence,
the acetylation and phosphorylation of choline sometimes compete for available substrate (Farber et al.,
1996; Ulus et al., 2006): When cholinergic neurons are forced to ﬁre frequently and to sustain the release of
ACh, choline’s incorporation into PC decreases (Farber et al., 1996) and the breakdown of membrane PC
increases (‘‘autocannibalism’’), liberating additional choline for ACh synthesis (Maire and Wurtman, 1985;
Blusztajn et al., 1986; Ulus et al., 1989). When the utilization of choline to form PC is increased (by
providing supplemental uridine and an omega‐3 fatty acid; see later), ACh synthesis is not diminished,
probably because so little choline is used for phosphatide formation compared with the amount used for
ACh synthesis (Ulus et al., 2006).
5.1 Biosynthesis of Acetylcholine
ACh is synthesized in cholinergic neurons—principally their terminals—by the ChAT‐mediated acetyla-
tion of free choline. Since, as described earlier, ChAT’s afﬁnity for choline is low compared with brain
choline levels, local choline concentrations normally control the rate of ACh synthesis (Blusztajn and
Wurtman, 1983), and treatments which increase brain choline (e.g., administering choline; Cohen and
Wurtman, 1975) or PC (Magil et al., 1981), or consuming choline‐rich foods (Cohen and Wurtman, 1976)
24 3.2 Choline and its products acetylcholine and phosphatidylcholine
rapidly cause parallel changes in brain ACh levels; in the amounts of ACh released when neurons ﬁre (Maire
and Wurtman, 1985; Jackson et al., 1995); and in postsynaptic ACh‐dependent functions like the control of
rat striatal (Ulus and Wurtman, 1976) and adrenomedullary (Ulus et al., 1977a, b, c) tyrosine hydroxylase
activities. The afﬁnity of ChAT for its other substrate, acetyl‐CoA—formed from glucose in mitochon-
dria—is substantially greater (K
¼77 mM; Rossier, 1977) than that for choline (K
¼540 mM), however
actual acetyl‐CoA concentrations in the vicinity of ChAT may still be insufﬁcient to saturate the enzyme,
and thus might also affect the rate of ACh synthesis. In support of this possibility, administration of glucose
has been found to stimulate ACh synthesis (Dolezal and Tucek, 1982), and to attenutate the depletion of
brain ACh induced by giving a muscarinic antagonist (Ricny et al., 1992). In microdialysis studies, glucose
enhanced the rise in ACh output produced by scopolamine (Ragozzino et al., 1994; Ragozzino and Gold,
1995). Systemic administration of glucose also increased hippocampal ACh release (Ragozzino et al., 1996,
1998; Kopf et al., 2001).
If choline levels in nerve terminals are reduced pharmacologically by administering a drug, HC3 that
blocks the reuptake of free choline from the synapse, the synthesis and release of ACh also decline in
parallel (Maire and Wurtman, 1985). Although such experiments conﬁrm the importance of choline
availability in controlling ACh synthesis, they do not necessarily allow it to be concluded that high‐afﬁnity
choline uptake is the rate‐limiting factor controlling intracellular choline levels or ACh biosynthesis. This
synthesis is affected by any process that modiﬁes the neuron’s concentration of free choline, and these levels
vary considerably as a function of plasma choline concentrations in addition, possibly, to changes in
reuptake efﬁciency. Moreover, the choline that enters the neuron via high‐afﬁnity uptake apparently is not
selectively used for acetylation as opposed to phosphorylation (Kessler and Marchbanks, 1979; Jope and
Jenden, 1981). As discussed earlier, it is possible, but not yet clearly demonstrated, that the density or
activity of high‐afﬁnity choline uptake sites in presynaptic membranes is affected by phosphorylation,
neuronal ﬁring, or the rate at which ACh is being released (Simon and Kuhar, 1975; Ferguson et al., 2003;
Gates et al., 2004).
5.2 Biosynthesis of Choline‐Containing Phosphatides
All cells use choline as an essential component of phospholipid subunits which, when aggregated, form all
of their membranes. The principal subunit, the phosphatide PC, is synthesized from choline by the CDP‐
choline cycle (or ‘‘Kennedy Cycle’’; Kennedy and Weiss, 1956) (>Figure 3.2‐4); PC, in turn, provides
the phosphocholine moiety for the synthesis of SM, the other major choline‐containing phospholipid
The CDP‐choline cycle involves three sequential enzymatic reactions (>Figure 3.2‐4): In the ﬁrst
(described earlier), catalyzed by CK, a monophosphate is transferred from ATP to the hydroxyl oxygen of
the choline, yielding phosphocholine. The second, catalyzed by CTP:phosphocholine cytidylyltransferase
(CT), transfers cytidylylmonophosphate (CMP) from CTP to the phosphorus of phosphocholine, yielding
cytidylyldiphosphocholine (CDP‐choline). The third and last reaction, catalyzed by CDP‐choline:1,2‐DAG
choline phosphotransferase (CPT), bonds the phosphocholine of CDP‐choline to the hydroxyl group on
the 3‐carbon of DAG, yielding the PC.
All these steps use compounds that the brain must obtain entirely or in part from the circulation, i.e.,
choline; a pyrimidine‐like uridine for conversion to CTP; a poluunsaturated fatty acid‐like DHA, and all
three steps can also affect the overall rate of PC synthesis in brain (Cansev et al., 2005; Wurtman et al.,
2006). Thus, choline administration increases brain phosphocholine levels in rats (Millington and
Wurtman, 1982) and humans (Babb et al., 2004), because CK’s K
for choline (2.6 mM; Spanner
and Ansell, 1979) is much higher than usual brain choline levels (35–60 mM). Most commonly, the second
CT‐catalyzed reaction is most rate limiting, either because not all of the CT is fully activated by being
attached to a cellular membrane (Vance and Pelech, 1984) or because local CTP concentrations are
insufﬁcient to saturate the CT (Ross et al., 1997). As described later, when brain CTP levels are increased
by giving animals uridine, CTP’s circulating precursor in humans (Cansev et al., 2005), PC synthesis is
Choline and its products acetylcholine and phosphatidylcholine 3.2 25
The activity of CPT and the extent to which this enzyme is saturated with DAG can also control the
overall rate of PC synthesis, for example, in PC12 cells extending neurites after exposure to the nerve growth
factor (NGF) (Araki and Wurtman, 1997). If rodents are given a diet that contains both choline and uridine
(as its monophosphate, UMP) and, by gavage, a PUFA (particularly the omega‐3 fatty acids DHA or EPA),
brain PC synthesis rapidly increases (Cansev et al., 2006; Wurtman et al., 2006), and absolute levels of PC
per cell (DNA) or per mg protein increase substantially (e.g., by 40–50% after 4 weeks of daily treatment
(Wurtman et al., 2006; >Table 3.2‐5). These treatments produce parallel or greater increases in each of
the other membrane phosphatides, as well as in proteins localized within synaptic membranes, like
synapsin‐1, PSD‐95 (>Figure 3.2‐7), and syntaxin‐3 (Fujita and Kurachi, 2000; Ferreira and Rapoport,
2002; Darios and Davletov, 2006). They also increase formation of dendritic spines (Sakamoto and Wurt-
man, 2006) and the release of ACh (Wang et al., 2004) and dopamine (Wang et al., 2005a) from striatal
neurons, and improve cognitive behaviors in aged rats (Teather and Wurtman, 2003), rats reared in a
socially deprived environment, or normal animals (Teather and Wurtman, 2005). Hence, the production,
Effects of UMP‐supplemented diet and/or DHA on brain phospholipid levels in gerbils
Treatments Total PL PC PE SM PS PI
Control diet þVehicle 403 23 155 8693473341202
UMP þDHA 436 15 188 8
UMP þDHA 502 12
Control diet þVehicle 351 8 152 6654452333212
UMP diet þVehicle 367 22 171 8
Control diet þDHA 392 20 185 12
UMP diet þDHA 442 24
Note: Groups of eight gerbils were given either a control diet and DHA’s vehicle (5% gum Arabic solution, by gavage) or a
UMP‐containing (0.5%) diet and DHA (300 mg/kg; in 5% gum Arabic solution, by gavage) for 1 or 3 weeks. In another set of
experiments, groups of eight gerbils were given either a control or a UMP‐containing (0.5%) diet, and received orally (by
gavage) DHA (300 mg/kg; in 5% gum Arabic solution) or just its vehicle for 4 weeks. At the end of each supplementation
period, the gerbils’ brains were harvested and assayed for phospholipids. Data are presented as nmol/mg protein. Data
from 1‐and 3‐week‐treated control diet and vehicle groups were pooled as there were no signiﬁcant differences among
P<0.001 when compared with data from control diet and vehicle group
Abbreviations: Total Pl, Total phospholipids; PC, phosphatidylcholine; PE, phosphatidylethanolamine; SM, sphingomyelin;
PS, phosphatidylserine; PI, phosphatidylinositol (Reproduced from Wurtman et al., 2006)
Increases in brain levels of PSD‐95 and Synapsin‐1 following dietary supplementation with uridine‐50‐mono-
phosphate (UMP) and docosahexaenoic acid (DHA). Groups of 8 gerbils were given either a control diet and
DHA’s vehicle (5% gum Arabic solution, by gavage) or a UMP‐containing (0.5%) diet and DHA (300 mg/kg; in
5% gum Arabic solution, by gavage) for 1 or 3 weeks. At the end of each supplementation period, the
gerbils’ brains were harvested and assayed for PSD‐95, Synapsin‐1, and beta‐tubulin. *P<0.05, **P<0.01,
and ***P<0.001 when compared with data from control diet and vehicle group (Reproduced from Wurtman
et al., 2006)
26 3.2 Choline and its products acetylcholine and phosphatidylcholine
.Figure 3.2‐7 (continued)
Choline and its products acetylcholine and phosphatidylcholine 3.2 27
levels, and functional properties of PC, and of other constituents of brain membranes, depend to a
surprising extent on blood levels of its three circulating precursors.
DAG molecules are highly heterogeneous with reference to their fatty acid constituents; however, those
containing DHA are preferentially used for phosphatide synthesis (Marszalek and Lodish, 2005; Marszalek
et al., 2005). The incorporation of circulating DHA to form these molecules involves several steps: the fatty
acid crosses the BBB (Hashimoto et al., 2002), then leaves the brain’s extracellular space by partitioning
into the external leaﬂet of a cell’s plasma membrane. It then ‘‘ﬂip‐ﬂops’’ to the inner leaﬂet, allowing it to
interact with intracellular fatty acid‐binding proteins and with the long‐chain fatty acyl‐CoA synthetase
(LCFAS; EC 184.108.40.206) to form an acyl‐fatty acid. This compound then attaches to the sn‐2 position of
glycerol‐3‐phosphate, a reaction catalyzed by acyl‐CoA synthetase long‐chain family member 6 (Acsl6;
Marszalek et al., 2005). This product then attaches a saturated fatty acid at the sn‐1 position to become
phosphatidic acid, which is subsequently dephosphorylated to form DAG. The concentration of DHA in
brain is only 1.2–2.9 mM (Deutsch et al., 1997; Contreras et al., 2000; Rosenberger et al., 2004) whereas the
of acyl‐CoA synthetase is an order of magnitude higher (26 mM; Reddy et al., 1984). Hence, the enzyme
is highly unsaturated with DHA, and very responsive to changes in DHA levels.
In vivo, orally administered DHA (200 mg/kg/day for 10 weeks to gerbils; Cao et al., 2006 or 300 mg/kg/
day for 12 weeks to rats; Hashimoto et al., 2002, 2005) has been shown to elevate brain DHA levels by
10–25%; however, this effect was not observed in gerbils (Cao et al., 2005) or rats (Boswell et al., 1996)
receiving 150 mg/kg/day or 1250 mg/kg/day, respectively, for 4 weeks, perhaps reﬂecting the slow mean turnover
time for brain PC, and the speed with which DHA‐containing phosphatides are deacylated and then reacylated
with less‐unsaturated fatty acids. In PC12 cells engineered to overexpress the Acsl6 protein, [
C]DHA added to
the medium is rapidly acylated to [
C]DHA‐CoA and incorporated into phospholipids and triglycerides
(Marszalek et al., 2005; Richardson and Wurtman, 2006). That exogenous DHA increases brain PC and PE
levels by increasing the phosphatide’s synthesis is further supported by the ﬁnding that DHA administration
concurrently raised PC and PE levels whereas lowering those of CDP‐choline and CDP‐ethanolamine
(Wurtman et al., 2006), the immediate precursors for PC and PE, with which the DHA‐containing DAG
presumably combined. DHA and other PUFA can also promote membrane synthesis by an additional
mechanism, i.e., by acting as agonists for a receptor, the syntaxin‐3 protein (Darios and Davletov, 2006).
As with DHA, the mechanisms by which supplemental uridine increases phosphatide synthesis proba-
bly include two processes, increased substrate saturation of the enzymes that convert uridine, via uridine‐50‐
triphosphate (UTP) and CTP, to endogenous CDP‐choline (Cansev et al., 2005), and activation by UTP of
speciﬁc G‐protein‐coupled receptors, the P2Y receptors which affect neurite outgrowth (Pooler et al., 2005).
A single dose of uridine, given as uridine‐50‐monophosphate (UMP) to gerbils by gavage, has been shown to
cause sequential increases in blood and brain uridine, then in brain UTP, CTP, and CDP‐choline levels
(Cansev et al., 2005). The uridine is carried across the BBB by a high‐afﬁnity concentrative nucleoside
transport protein, CNT2, described later (Li et al., 2001; Gray et al., 2004); this protein’s afﬁnity for cytidine
is only one‐tenth or one‐twentieth that for uridine (Larrayoz et al., 2006; Nagai et al., 2006a, b). Moreover,
in contrast to uridine, little or no cytidine is present in human blood, any exogenous or endogenous
circulating cytidine being rapidly converted to uridine in the human liver (Wurtman et al., 2000).
The effect of the uridine on phosphatide synthesis is short‐lived, and insufﬁcient in itself to produce
reliable increases in brain PC or PE. However, chronic consumption of a uridine source (via the diet) for
three or four weeks raises brain PC and PE levels substantially (>Table 3.2‐5), even though the increases
that this treatment produces in plasma uridine are small and transient. In vitro, uridine in concentrations of
50 mM or greater increases the size of neurites (Pooler et al., 2005) sprouting from NGF‐stimulated PC12
cells (Araki and Wurtman, 1997); it also accelerates phosphatide synthesis (Richardson et al., 2003) and
increases neuroﬁlament protein levels in these cells (Pooler et al., 2005). The stimulatory effect of uridine on
neuritogenesis also involves activation by UTP, its product, of P2Y receptors. This was shown by demon-
strating that a lower concentration of UTP (10 mM) than that of free uridine can enhance neurite outgrowth
(Pooler et al., 2005), and that the effects of both uridine and UTP on neuritogenesis are blocked by apyrase,
a drug that degrades nucleotides, or by P2Y antagonists (Pooler et al., 2005).
28 3.2 Choline and its products acetylcholine and phosphatidylcholine
Combined treatment of animals with dietary choline, UMP, and DHA tends to produce larger increases
in levels of individual brain phosphatides (e.g., SM) than the sum of the increases caused by the individual
precursors (>Table 3.2‐5). This probably reﬂects the operation of all the earlier mechanisms: Providing
each of the three circulating precursors decreases the likelihood that its intracellular levels will become
limiting when levels of the other precursors have been increased, and providing uridine or DHA enhances
the activation of P2Y receptors by UTP, or of syntaxin‐3 by the DHA. The relevant P2Y receptors might
include any of the pyrimidine‐sensitive receptors (P2Y2, P2Y4, or P2Y6) in the P2Y family.
Giving the three precursors concurrently increased PC by 21% (nmol/mg protein) after 1 week, and by
40% or 45% after 3 or 4 weeks; corresponding increases expressed as nmol/mg DNA were by 20%, 34%, and
41%, respectively (>Table 3.2‐5). Similar or greater changes are noted in the other phosphatides, or, in rat
pups if the precursors were administered to their mothers during pregnancy and lactation (Marzloff et al.,
2006). It is conceivable that even greater increases might be produced by administering larger doses of the
precursors, or by extending the treatment for longer periods. The amounts of choline provided in the
studies cited (Marzloff et al., 2006; Wurtman et al., 2006) were standard for gerbil and rat chows; UMP,
the uridine source, is not normally included in such chows; however, it is present in human mother’s milk
and included in infant formulas in amounts intended to provide up to 0.3 mg/100 J (Yu, 2002); DHA is not
a constituent of gerbil or rat chow, but is variably present in human diet depending in large part on
the quantities of seafoods—particularly higher‐fat ﬁsh like herring and salmon (Harris, 2005)—that a
person elects to eat. Consumption of up to 3 g/day of DHA and other omega‐3 fatty acids (e.g.,
eicosapentaenoic acid) is generally regarded as safe (GRAS) by the U. S. Food and Drug Administration,
and doses of 4 g/day, or 50–60 mg/kg are sometimes used in the treatment of hypertriglyceridemia
(Stalenhoef et al., 2000). The precise ratios of supplemental choline:uridine:DHA needed for optimal
enhancement of brain phosphatide levels await determination, as does the question of whether AA or
other PUFA present in phosphatides besides DHA should also be included. No data are presently available
on possible behavioral consequences of increasing the amounts of synaptic membrane in human brain by
administering DHA, a uridine source, and choline. Brain levels of choline, ethanolamine (Nitsch et al.,
1992), and DHA (Soderberg et al., 1991) are all known to be subnormal among patients with Alzheimer’s
disease, and brains of such patients exhibit characteristic decreases in synaptic size and number (Terry et al.,
1991; Selkoe, 2002; Coleman et al., 2004), as well as increases in the phosphatide breakdown products GPC
and glycerophosphoethanolamine (Blusztajn et al., 1990; Nitsch et al., 1992). If brains of patients with
Alzheimer’s disease remain capable of responding to the three circulating phosphatide precursors by
increasing brain levels of synaptic membrane, the precursors could conceivably confer some therapeutic
beneﬁt in this disease.
5.2.1 CTP:Phosphocholine Cytidylyltransferase
CTP:phosphocholine cytidylyltransferase (CT; EC 220.127.116.11) catalyzes the condensation of CTP and phos-
phocholine to form CDP‐choline (>Figure 3.2‐4). CT is present in both soluble and particulate fractions of
the cell (Wilgram and Kennedy, 1963); the cytosolic form is reportedly inactive whereas the membrane‐
bound form is active (Vance and Pelech, 1984; Tronchere et al., 1994). Increases in the association of CT
with membranes reportedly correlate with increases in CT activity and in the net synthesis of PC in vitro
(Sleight and Kent, 1980, 1983; Pelech et al., 1984). Some other lipids (reviewed in Cornell and Northwood,
2000) and DAG (Sleight and Kent, 1980; Utal et al., 1991) can stimulate the translocation of CT from the
cytosol to membranes in vitro, thereby activating the enzyme. However translocation may not be the sole
mechanism of CT activation, inasmuch as increases in the activity of membrane‐bound CT do not always
correlate with decreases in that of the cytosolic enzyme (Weinhold et al., 1991); as would be expected
if traslocation were the only means whereby CT become activated. The phosphorylation state of CT may
also be important (Watkins and Kent, 1991) as well as its substrate saturation with CTP and perhaps
Choline and its products acetylcholine and phosphatidylcholine 3.2 29
CT has been puriﬁed to homogeneity (Weinhold et al., 1986), and has been cloned from rat liver
(Kalmar et al., 1990) and from a human erythroleukemic cell line (Kalmar et al., 1994). The puriﬁed form
exists as an elongated dimer (Cornell, 1989). Mammalian CT proteins are divided into four functional
domains: an N‐terminal nuclear targeting sequence, a catalytic domain, a membrane–lipid binding
domain, and a C‐terminal phosphorylation domain. CT is termed CTaor CTbdepending on whether or
not the N‐terminal nuclear targeting domain does or does not contain a nuclear localization signal. The two
genes that encode CT proteins reside on distinct chromosomes, i.e., Pcyt1a, which encodes the CTaisoform
and is located on human chromosome 3q (Tang et al., 1997; Karim et al., 2003), and Pcyt1b, which encodes
CTb1, ‐b2, and –b3 isoforms and is X‐linked (Lykidis et al., 1998, 1999; Karim et al., 2003).
s of CT for CTP and phosphocholine in brains of laboratory rodents and humans are 1–1.3 mM
and 0.30–0.31 mM (Mages et al., 1988; Ross et al., 1997), respectively, whereas brain levels of these
compounds are 70–110 mM (Mandel and Edel‐Harth, 1966; Abe et al., 1987; Cansev et al., 2005) and
0.32–0.69 mM (Millington and Wurtman, 1982; Nitsch et al., 1992; Klein et al., 1993), respectively. Hence,
brain CT is normally unsaturated with both of its substrates but especially CTP, suggesting a limiting role
for cellular CTP in PC synthesis. In fact, treatments that increase cellular CTP levels do enhance the
synthesis of CDP‐choline and PC in poliovirus‐infected HeLa cells (Choy et al., 1980); undifferentiated
PC12 cells (Lopez G‐Coviella and Wurtman, 1992; Richardson et al., 2003); rat striatal brain slices (Savci
and Wurtman, 1995); and gerbil brain in vivo (Cansev et al., 2005).
5.2.2 CDP‐Choline:1,2‐Diacylglycerol Cholinephosphotransferase
CDP‐choline:1,2‐DAG cholinephosphotransferase (CPT; EC 18.104.22.168) catalyzes the ﬁnal reaction in the
Kennedy cycle; it transfers the phosphocholine moiety from CDP‐choline to DAG, thus yielding PC and
releasing CMP (>Figure 3.2‐4). CPT, an integral membrane protein, is present primarily in the endoplas-
mic reticulum (Coleman and Bell, 1977). The enzyme protein has been solubilized and partially puriﬁed
from microsomes of rat liver (Kanoh and Ohno, 1976; Ishidate et al., 1993), rat brain (Roberti et al., 1989),
and hamster liver (O and Choy, 1990); puriﬁcation to homogeneity has not been accomplished as the
enzyme is membrane‐bound, and thus is susceptible to detergents. A human cDNA has been isolated which
codes for an enzyme with both cholinephosphotransferase and ethanolaminephosphotransferase (EPT)
activities (hCEPT1; Henneberry and McMaster, 1999), and a different human cDNA has also been isolated
which codes for an enzyme exhibiting only cholinephosphotransferase‐speciﬁc activit y (hCPT1; Henneberry
et al., 2000). CPT may be a reversible enzy me, synthesizing CDP‐choline from PC and CMP in liver (Kanoh
and Ohno, 1973a, b) or brain microsomal preparations (Goracci et al., 1981, 1986; Roberti et al., 1992).
The CPT reaction is unsaturated with the enzyme’s substrates; its K
values for CDP‐choline and DAG
in rat liver are 200 mM and 150 mM (Cornell, 1992) respectively, whereas the concentrations of these
compounds in liver are 40 mM (Korniat and Beeler, 1975) and 300 mM (Turinsky et al., 1991), respectively
(A DAG concentration of at least 1000 mM would probably be needed to stimulate the enzyme). Brain CDP‐
choline and DAG levels are even lower, i.e., about 10–30 mM (Alberghina et al., 1981; Cansev et al., 2005)
and 75 mM (Abe et al., 1987), respectively. Levels of cellular DAG have been shown to limit PC synthesis
in permeabilized HeLa cells (Lim et al., 1986), cultured rat hepatocytes (Jamil et al., 1992), and PC12
cells (Araki and Wurtman, 1997). In the latter study, cellular DAG levels had been increased by ﬁvefold or
none by exposing them to NGF. This treatment differentiated the cells, causing neurite outgrowth and a
major increase in PC levels. None of these studies distinguished between the enzymes that act on both
choline and ethanolamine (PECT1) and the enzyme that acts only on choline (PCT1). A more recent
report, using cloning and expression methods, described that the K
of human PECT1 for CDP‐choline, as
36 mM (Wright and McMaster, 2002), which would probably still be too high to be saturated with this
substrate in brain. The K
of the enzyme for its substrates might also be affected by the fatty
acid composition of the DAG molecule; for example, incubating mouse liver microsomes with DAG
molecules that contained two oleic acids (1,2‐dioleoyl‐sn‐glycerol; Di‐C
(cis‐9)) rather than two palmitic
acids (1,2‐dipalmitoyl‐sn‐glycerol [Di‐C
]), increased its K
s for DAG from 86 6mM to 1860 39 mM
30 3.2 Choline and its products acetylcholine and phosphatidylcholine
and its K
for CDP‐choline from 41 2mM to 1000 141 mM (Mantel et al., 1993). Hence the enzyme’s
afﬁnity for its substrates declined by 20–25‐fold.
5.2.3 Uptake of Uridine and Cytidine into Brain Cells
Uridine and cytidine are transported across cell membranes, including the BBB, through two families of
transport proteins, i.e., the Na
‐independent, low‐afﬁnity, equilibrative transporters (ENT1 and ENT2)
and the Na
‐dependent, high‐afﬁnity, concentrative (CNT1, CNT2, and CNT3) nucleoside transporters
(Baldwin et al., 2004; Gray et al., 2004). The two ENT proteins, which transport uridine and cytidine with
similar afﬁnities, have been cloned from rat (Redzic et al., 2005) and mouse (Murakami et al., 2005) BBB.
Inasmuch as their K
values for the pyrimidines are in the high micromolar range (100–800 mM; Pastor‐
Anglada et al., 1998) they probably mediate BBB pyrimidine uptake only when plasma levels of uridine and
cytidine have been elevated experimentally. In contrast, CNT2, which transports both uridine and purines
like adenosine probably mediates uridine transport across the BBB under physiologic conditions. The K
values for the binding of uridine and adenosine to this protein (which has been cloned from rat BBB; Li
et al., 2001) are in the low micromolar range (9–40 mM in kidney, intestine, spleen, liver, macrophage, and
monocytes; Grifﬁth and Jarvis, 1996), whereas plasma uridine levels are subsaturating, i.e., 0.9–3.9 mMin
rats (Traut, 1994); 3.1–4.9 mM in humans (Traut, 1994); and around 6.5 mM in gerbils (Cansev et al., 2005).
Cytidine has not been thought to be a substrate for CNT2 (Gray et al., 2004); however, recent studies
suggest that CNT2 can also transport it, but with a much lower afﬁnity than that for uridine (Larrayoz et al.,
2006; Nagai et al., 2006a, b).
Like other circulating compounds, pyrimidines may also be taken up into brain through the epithelium
of the CP and the ENT1, ENT2, and CNT3 transporters (Baldwin et al., 2004; Gray et al., 2004); all these
proteins have been found in CP epithelial cells of rats (Anderson et al., 1999a, b; Redzic et al., 2005) and
rabbits (Wu et al., 1992, 1994). However, the surface area of BBB is probably 1000 times that of the CP
epithelium (i.e., in humans 21.6 m
vs. 0.021 m
; Pardridge, 2001); hence, in any event, the BBB is the major
locus at which circulating uridine enters the brain. These more recent data on speciﬁc transport proteins
conﬁrm those from earlier studies on pyrimidine uptake across the BBB, which ﬁrst demonstrated uridine’s
efﬁcient (Hogans et al., 1971; Cornford and Oldendorf, 1975) and cytidine’s poor (Galletti et al., 1991)
uptake into brain. Moreover, cytidine is barely measurable in human plasma (Traut, 1994; Wurtman et al.,
2000), and, as described later, when exogenous cytidine is given in the form of CDP‐choline, it appears in
human blood as uridine, not cytidine (Wurtman et al., 2000). Hence, uridine is the preferred circulating
precursor for the CTP used in humans and some rodents (e.g., gerbils but not rats) for brain phosphatide
synthesis (Cansev and Wurtman, 2005; Cansev, 2006).
5.2.4 Phosphorylation of Uridine and Cytidine to UTP and CTP
Uridine and cytidine are converted to their respective nucleotides following successive phosphorylations by
various kinases (>Figure 3.2‐4). Uridine‐cytidine kinase (UCK) (ATP:uridine‐50‐phosphotransferase, EC
22.214.171.124), the ﬁrst enzyme in this cascade, catalyzes the phosphorylations of uridine and cytidine to form
UMP and cytidine‐50‐monophosphate (CMP), respectively (Canellakis, 1957; Skold, 1960; Orengo, 1969).
Several different forms of UCK exist, possibly as isoenzymes (Krystal and Webb, 1971; Absil et al., 1980).
Humans have two such isoenzymes, UCK1 and UCK2, which have now been cloned (Koizumi et al., 2001;
van Rompay et al., 2001).
The next enzyme in this sequence, which phosphorylates UMP and CMP to form uridine‐50‐diphos-
phate (UDP) and CDP, respectively, is UMP–CMPK (ATP:CMP phosphotransferase, EC 126.96.36.199)
(Hurwitz, 1959; Sugino et al., 1966; Ruffner and Anderson, 1969). UDP and CDP are further phosphory-
lated to UTP and CTP, by nucleoside diphosphate kinases (NDPK) (Nucleoside triphosphate:Nucleoside
diphosphate phosphotransferase, EC 188.8.131.52) (Berg and Joklik, 1954; Parks and Agarwal, 1973). The
Choline and its products acetylcholine and phosphatidylcholine 3.2 31
mRNAs for UCK1 (van Rompay et al., 2001) and UMP–CMPK (van Rompay et al., 1999) as well as NDPK
activity have been described in brain (Langen et al., 1999; Kim et al., 2002).
The interconversions of uridine and cytidine, and of their respective nucleotides, are also observed in
mammalian cells. Cytidine and CMP can be deaminated to uridine and UMP (Wang et al., 1950), whereas
UTP is aminated to CTP by CTP synthase [UTP:ammonia ligase (ADP‐forming), E.C. 184.108.40.206] (Lieberman,
1956; Hurlbert and Kammen, 1960). This enzyme acts by transferring an amide nitrogen from glutamine to
the C‐4 position of UTP, thus forming CTP (Zalkin, 1985). CTP synthase activity has been demonstrated in
rat brain (Genchev and Mandel, 1974).
All the enzymes described earlier apparently are unsaturated with their respective nucleosides or
nucleotides in brain and other tissues. For example, K
s for uridine and cytidine of UCK prepared from
various tissues varied between 33 and 270 mM (Skold, 1960; Orengo, 1969; Anderson, 1973; Greenberg
et al., 1977), and the K
for uridine of recombinant enzyme cloned from mouse brain was 40 mM (Ropp
and Traut, 1996, 1998). Brain uridine and cytidine levels are about 22–46 pmol/mg wet weight (Mascia
et al., 1999; Cansev et al., 2005) and 6–43 pmol/mg wet weight (Peters et al., 1987; Cansev et al., 2005),
respectively. Hence, the syntheses of UTPand CTP, and the subsequent syntheses of brain PC and PE via the
Kennedy pathway, depend on the availability of their pyrimidine substrates. Indeed, an increase in the
supply of uridine or cytidine to neuronal cells, in vitro (Savci and Wurtman, 1995; Richardson et al., 2003;
Pooler et al., 2005) or in vivo (Cansev et al., 2005; Cansev and Wurtman, 2005), enhanced the phosphory-
lation of uridine and cytidine, elevating the levels of UTP, CTP, and CDP‐choline.
6 Physiological and Behavioral Effects of Choline
Administration of choline by direct placement into the CNS, orally or by injection, can produce numerous
physiological or behavioral effects. Some of these are readily attributable to enhanced ACh release; others
may be mediated by phospholipid metabolism or, conceivably, by direct actions of the choline on
cholinergic receptors, as discussed earlier. Some remain unexplained.
6.1 Blood Pressure
Intravenous choline administration lowers blood pressure in both humans and animals (Mendel et al.,
1912; Steigmann et al., 1952; Anton, 1954; Kapp et al., 1970; Singh, 1973; Savci et al., 2003). Intramuscular
administration of choline (20 mg/kg/day for 3 days) to rats attenuates the fall in blood pressure induced by
acute hemorrhage, and increases survival rate (Altura, 1978). Intraperitoneal choline (60 mg/kg) partially
restores blood pressure after the induction of hypotension by acute hemorrhage (Ulus et al., 1995); in
contrast intravenous choline (54 mg/kg) further decreases blood pressure and can cause death in hemor-
rhaged rats (Savci et al., 2003). Oral choline fails to affect cardiovascular function in rats (Ulus et al., 1979)
but reportedly lowered blood pressure slightly in some patients with Alzheimer’s disease (Boyd et al., 1977).
Intracerebroventricular choline (8 mg/dog) produced a biphasic blood pressure response in anesthe-
tized dogs, an immediate and short‐lasting (about 10–15 min) blood pressure rise (by 60–90 mm Hg)
followed by a longer‐lasting (about 60 min) fall (by 15–20 mm Hg; Srimal et al., 1969). In rats, intracisternal
choline (12.5–50 mg; Kubo and Misu, 1981a) or its microinjection (1–3 mg/site; Kubo and Misu, 1981b) into
the dorsal medulla lowered blood pressure, whereas intracerebroventicular choline (50–150 mg/rat) raised
blood pressure and decreased heart rate (Caputi and Brezenoff, 1980; Arslan et al., 1991; Isbil‐Buyukcoskun
et al., 2001; Li and Buccafusco, 2004) for 5–20 min. In rats, intracerebreventricular choline (25–150 mg/rat)
restored normal blood pressure among animals made hypotensive by acute hemorrhage (Ulus et al., 1995;
Savci et al., 2002b), endotoxin (Savci and Ulus, 1997), chemical sympathectomy (Gurun et al., 1997a),
spinal cord transection (Savci and Ulus, 1998), autonomic ganglion blockade, or a‐adrenoceptor blockade
(Savci and Ulus, 1996). At a dose of 180 nmol, choline also potentiated the pressor responses evoked by
naloxone or glycyl‐glutamine (Gurun et al., 2003). In normotensive rats, the blood pressure responses to
choline administered centrally involve local activation of both mAChRs and nAChRs (Arslan et al., 1991);
32 3.2 Choline and its products acetylcholine and phosphatidylcholine
including the a7 nAChR subtype (Li and Buccafusco, 2004). In hypotensive animals, these blood pressure
responses involve presynaptic activation of central nAChRs (Ulus et al., 1995; Savci and Ulus, 1996, 1997,
1998; Gurun et al., 1997a; Savci et al., 2002b).
6.2 Body Temperature
Intracerebroventricular (75–300 mg/rat), but not intraperitoneal (30–120 mg/kg), choline decreases body
temperature (Unal et al., 1998). This hypothermia is mainly mediated by M1and M3 muscarinic receptors
(Unal et al., 1998).
Choline can alter responses to painful stimuli in experimental animals, and can modify the actions of
analgesic drugs. Given intraperitoneally (15–60 mg/kg) to rats, it diminishes the analgesic actions of
morphine (10 mg/kg; subcutaneously) as assessed using the hot‐plate test (Botticelli et al., 1977). In
mice, subcutaneous (up to 500 mg/kg; Damaj et al., 2000) or intravenous (2–64 mg/kg; Wang et al.,
2005b) choline fails to alter responses to thermal pain, but when given intracerebroventricularly (Damaj
et al., 2000; Wang et al., 2005b) or intrathecally (Damaj et al., 2000) at doses of 30–120 mg/animal, it
produces antinociception in the same pain model. This latter effect can be blocked by the nonselective
mAChRs antagonist atropine or by antagonists of a7 nAChRs (e.g., metillycaconitine; a‐bungarotoxin).
Intravenous choline (4–64 mg/kg) produced signiﬁcant antinociception in mice in the late phase of
inﬂammatory pain responses to subcutaneous injection of 5% formalin (Wang et al., 2005b). At a 2 mg/kg
dose, it also enhanced the antinociceptive effects of aspirin (9.4 mg/kg; i.v.) and morphine (0.165 mg/kg; i.v.).
This action was also blocked by the a7 nAChR antagonists metillycaconitine and a‐bungarotoxin, but not by
atropine or naloxone (Wang et al., 2005b).
6.4 Neuroendocrine Effects
Choline produces a variety of neuroendocrine responses when administered peripherally or centrally to
rats. Its oral administration (20 mmol/kg by stomach tube) for 4 days increased urinary catecholamine
output (Scally et al., 1978). Given intraperitoneally, choline (30–120 mg/kg) elevated plasma catechola-
mines (Ilcol et al., 2002c) and insulin (Ilcol et al., 2003a). Given intracerebroventricularly (50–150 mg/rat),
it increased plasma concentrations of the catecholamines (Arslan et al., 1991; Ulus et al., 1995; Gurun et al.,
2002); vasopressin (Arslan et al., 1991; Ulus et al., 1995; Savci and Ulus, 1996, 1998; Gurun et al., 1997a;
Savci et al., 2003); ACTH (Savci et al., 1996); bendorphin (Savci et al., 1996); and prolactin (Gurun et al.,
1997b). The increases in plasma vasopressin, ACTH, and b‐endorphin were found to be mediated by central
nAChRs, and that of prolactin by mAChRs (Gurun et al., 1997b). The increases in plasma insulin after
intraperitoneal choline involve both ganglionic nAChRs and the M1 and M3 subtypes of mAChRs (Ilcol
et al., 2003a). Choline’s site of action in producing rest of these neuroendocrine effects is principally
presynaptic, i.e., via enhancing ACh release (Scally et al., 1978; Savci et al., 1996, 1998, 2003; Ilcol et al.,
2003a), and the effects are enhanced by hemorrhagic, hypotensive, or osmotic stresses (Scally et al., 1978;
Ulus et al., 1995; Savci et al., 1996, 1998, 2003; Ilcol et al., 2003a).
6.5 Peripheral Metabolism
Peripheral (40–120 mg/kg; intraperitoneal) or central (75–300 mg/rat; icv) administration of choline
increases blood glucose levels in rats (Gurun et al., 2002; Ilcol et al., 2002c). This hyperglycemic response
is prevented by blockade of ganglionic nAChRs or a‐adrenoceptors, as well as by bilateral adrenalectomy
Choline and its products acetylcholine and phosphatidylcholine 3.2 33
(Ilcol et al., 2002c), and apparently is mediated by the stimulation of adrenomedullary catecholamine
release and subsequent activation of a‐adrenoceptors. It apparently also involves activation of central
nAChRs affecting the sympathoadrenal system (Gurun et al., 2002). As decribed earlier, glucose is a source
of acetyl‐CoA, which, like choline, is a limiting precursor for ACh. Hence, choline administration may
enhance ACh synthesis by this additional mechanism.
Acute administration of choline to humans, as choline chloride or as PC, improved short‐term memory in
some studies (Leathwood et al., 1982; Ladd et al., 1993) but not in others (Davis et al., 1980; Harris et al.,
1983). Intraperitoneal choline (6–60 mg/kg) in combination with glucose (10–30 mg/kg) improved passive
avoidance behavior in mice (Kopf et al., 2001). Chronic consumption of a choline‐rich diet by aged mice
reportedly counteracted the age‐associated decline in learning and memory (Bartus et al., 1980; Golczewski
et al., 1982; Leathwood et al., 1982).
Supplementation of pregnant or lactating rats with choline chloride during the perinatal period
(embryonic days 12–17 and/or postnatally 16–30 days) caused long‐lasting improvements in spatial
memory, as illustrated using radial‐arm maze tests (Meck et al., 1988, 1989; Meck and Williams, 1997b,
1999; Williams et al., 1998) and the Morris water maze test (Schenk and Brandner, 1995; Tees, 1999a, b; Tees
and Mohammadi, 1999). In adulthood, rats that had received supplemental choline (about four times
dietary levels) perinatally exhibited increased memory capacity (Meck and Williams, 1997c) and precision
(Meck and Williams, 1999), and performed more accurately on tests of spatial memory (Meck et al., 1988,
1989; Meck and Williams, 1997a, b, c, 1999; Williams et al., 1998). Moreover, perinatal supplementation
with choline provided some protection against memory impairments usually associated with normal aging
(Tees and Mohammadi, 1999), neonatal alcohol exposure (Thomas et al., 2004; Wagner and Hunt, 2006),
or epileptic seizures (Yang et al., 2000; Holmes et al., 2002).
Although the mechanism by which perinatal choline supplementation produces these long‐term effects
on memory remains unclear, such supplementation is known also to cause enduring neuroanatomic and
neurochemical changes in brain regions involved in memory. For example, choline supplementation
increased hippocampal ChAT activity, mAChRs density (Meck et al., 1989), ACh release (Cermak et al.,
1998), and the amplitude of ACh‐mediated excitatory potentials (Montoya et al., 2000), and it reduced
acetylcholinesterase activity (Cermak et al., 1998, 1999). Moreover, basal forebrain cholinergic neurons
projecting to the hippocampus were larger and more spherical among rats that had been supplemented
perinatally with choline (Loy et al., 1991; Williams et al., 1998). Prenatal choline supplementation also
reportedly enhanced MAPK and CREB activation (Mellott et al., 2004); N‐methyl‐D‐aspartate (NMDA)
receptor‐mediated neurotransmission (Montoya and Swartzwelder, 2000); long‐term potentiation (LPT)
(Pyapali et al., 1998); PLD activity (Holler et al., 1996); NGF levels (Sandstrom et al., 2002); and dendritic
spine formation (Mervis, 1982) in rat hippocampus. It also decreased the rate of apoptosis in the
hippocampus and basal forebrain of 18‐day old fetuses (Holmes‐McNary et al., 1997) and increased
brain cell division (Albright et al., 1999a, b). Prenatal supplementation with choline also was associated
with greater excitatory responsiveness, reduced slow afterhyperpolarizations, enhanced afterdepolarizing
potentials, larger somata, and greater basal dendritic arborization in hippocampal CA1 pyramidal cells
studied postnatally at 20–25 days of age (Li et al., 2003).
6.7 Drug Interactions
Acute or chronic choline administration can modify the actions of some centrally active drugs. Chronic
(28–35 days) treatment of rats with choline through the diet (about ten times more choline than in control
diets) produced behavioral hyperactivity and attenuated the sedative/hypnotic and hypotermic effects of
pentobarbital (Wecker et al., 1987). Chronic choline supplementation also increased the density of binding
sites for nicotine (Coutcher et al., 1992) and a‐bungarotoxin (Morley and Garner, 1986) and produced
34 3.2 Choline and its products acetylcholine and phosphatidylcholine
tolerance to the convulsive and lethal actions of nicotine (Wecker et al., 1982). Chronic dietary choline
supplementation to mice modulated benzodiazepine receptor binding and g‐aminobutyric acid receptor
function (Miller et al., 1989), also decreasing seizure activity and the lethality of such seizure‐promoting
drugs such as nicotine, paraoxon, strychnine, and pentylenetetrazol (Wecker et al., 1982). Acute intraperi-
toneal choline administration (100 mg/kg) to morphine‐dependent rats decreased withdrawal symptoms
and the associated weight loss (Pinsky et al., 1973; Frederickson and Pinsky, 1975).
6.8 Neuroprotective and Cytoprotective Effects
Choline exhibits cytoprotective and neuroprotective actions in vivo and in vitro: In vitro, choline itself or
some of its analogs (i.e., mono‐,di‐, and triethylcholine and pyrrolidinecholine), can, at 1–10 mM
concentrations, protect against manifestations of cytotoxicity in differentiated PC12 cells induced by
growth factor deprivation (Jonnala et al., 2003). Choline (5–75 mM) suppresses the development of dark
cell degeneration in Purkinje neurons following receptor activation with AMPA (DL‐amino‐3‐hydroxy‐5‐
methyl‐isoxazole‐4‐propionic acid; Strahlendorf et al., 2001). At concentrations of 1 mM or greater it also
reduces NMDA toxicity in organotypic hippocampal slice cultures derived from neonatal rats (Mulholland
et al., 2004), and reportedly it is cytoprotective in PC‐12 cells (Jonnala et al., 2003); Purkinje neurons
(Strahlendorf et al., 2001); and hippocampal cells (Mulholland et al., 2004)—an effect mediated by the a7
subtype of nAChRs. Prenatal choline supplementation for 6 days during days E12–E17 of gestation
protected against subsequent neurodegeneration in the posterior cingulate and retrosplenial cortices,
induced in female adolescent rats by peripheral administration of dizocilpine (Guo‐Ross et al., 2002).
In cultured neonatal cardiac ventricular cells, choline (0.1–1 mM) reduced H
‐induced apoptotic cell
death, by acting via M3 mAChRs (Yang et al., 2005). In vivo, choline (20 mg/kg; i.v.) attenuated endotoxin‐
induced multiple organ injury (i.e., renal, hepatic, and cardiac injuries) in dogs (Ilcol et al., 2005b).
Similarly, in rats, choline supplementation in the diet for 3 days attenuated endotoxin‐induced hepatic
injury and improved survival (Rivera et al., 1998). Choline‐induced protection from the tissue injuries
induced by endotoxin is associated with a reduction in serum levels of tumor necrosis factor‐a(TNF‐a)
(Rivera et al., 1998; Ilcol et al., 2005b) and with the improvement in platelet counts and platelet closure
times (Yilmaz et al., 2006a). Intravenous choline administration (5 mg/kg) protected rats (5 mg/kg; i.v.)
from ischemic myocardial injuries by stimulating M3‐mAChRs (Yang et al., 2005).
7 Effects of Exogenous CDP‐Choline
Cytidine‐50‐diphosphocholine (CDP‐choline; citicoline), which is composed of choline and cytidine linked
by a diphosphate bridge, is both an essential intermediate in the synthesis of endogenous PC through the
Kennedy cycle (>Figure 3.2‐4), and a drug used in some countries to treat cerebral ischemia, traumatic
brain injury, Parkinson’s disease, or stroke.
When administered orally or parenterally, exogenous CDP‐choline is completely hydrolyzed, ﬁrst to
cytidine monophosphate and phosphocholine, and then to free cytidine and choline (Lopez G‐Coviella
et al., 1987). In humans, the cytidine is further transformed to uridine, hence giving CDP‐choline to
humans causes dose‐related increases in serum uridine and choline levels but not in serum cytidine
(Wurtman et al., 2000; Cansev, 2006). In laboratory rodents, depending on species and on the activity of
the hepatic enzyme cytidine deaminase (Chabot et al., 1983; Ku
¨hn et al., 1993), which converts cytidine to
uridine, CDP‐choline administration can principally elevate either serum uridine (e.g., in gerbils) or
cytidine (e.g., in rats), besides choline. Moreover, as treatments that elevate plasma uridine (or cytidine)
and choline thereby increase brain PC synthesis (Wurtman et al., 2000; Richardson et al., 2003; Cansev
et al., 2005; Ulus et al., 2006) as well as steady‐state brain levels of PC and other membrane phosphatides
(Lopez G‐Coviella et al., 1992, 1995; Wurtman et al., 2006), it is likely that some of CDP‐choline’s
therapeutic actions result from changes that it produces in the quantities or the composition of brain
membranes. Some other effects of exogenous CDP‐choline (Savci et al., 2002a, 2003; Cavun and Savci, 2004;
Choline and its products acetylcholine and phosphatidylcholine 3.2 35
Cavun et al., 2004) probably are mediated by increasing AChrelease, secondary to the rise it produces in plasma
and brain choline levels (Savci et al., 2002a, 2003).
By increasing brain levels of endogenous CDP‐choline, exogenous CDP‐choline also increases the
amounts of DAG that combine with this intermediate to form PC (and that combine with endogenous
CDP‐ethanolamine to form PE) (Araki and Wurtman, 1998). CDP‐choline administration can also affect
the brain by increasing the amount of free AA that is used to form DAG, which then combines with
endogenous brain CDP‐choline. This causes a decrease in free AA levels, which might otherwise be
neurotoxic, and thus it decreases the ultimate size of the brain damage following a stroke and brain injury
(Warach et al., 2000). This reduction in AA may be the major mechanism underlying CDP‐choline’s acute
therapeutic effects (Lopez G‐Coviella et al., 1998). It has also been suggested that exogenous CDP‐choline
may decrease brain levels of free AA by directly inhibiting phospholipase A2 activity or by decreasing
the formation of that enzyme protein (Adibhatla and Hatcher, 2003; Adibhatla et al., 2006).
7.1 Hypoxia and Ischemia
Intracerebral CDP‐choline (0.6 mmoles) prevented the ischemia‐induced loss of radioactive choline from
glycerophospholipids, and suppressed the increase in brain levels of free fatty acids in a global model of
ischemia in rats (Dorman et al., 1983; Goldberg et al., 1985). In a rat model of transient cerebral ischemia,
intraperitoneal CDP‐choline (250 mg/kg; twice a day for 4 days) improved neurological signs and
attenuated the increases in glucose and pyruvate levels and the decrease in ACh synthesis from labeled
glucose (Kakihana et al., 1988). In an ischemic and anoxic rat model, CDP‐choline (300 mg/kg, i.p.)
decreased the incidence of neurological deﬁcits (Yamamoto et al., 1990). In a chronic hypoxia rat model
produced by placing animals in chambers in which the oxygen content was depressed (7–15%) for extended
time periods, CDP‐choline (100 mg/kg in food) protected vigilance behavior (Hamdorf and Cervos‐
Navarro, 1990), reduced hypoxia‐induced behavioral deterioration (Hamdorf and Cervos‐Navarro, 1991),
and increased survival time at 7% O
(Hamdorf and Cervos‐Navarro, 1991; Hamdorf et al., 1992). In a
model of rat experimental hypoxia induced by giving potassium cyanide, oral CDP‐choline given for 4 days
before the induction of hypoxia increased survival time (Tornos et al., 1983a). Araki et al. (1988) also
observed a neuroprotective effect of CDP‐choline in mice in which the cerebral ischemia was induced by
decapitation or by potassium cyanide intoxication. CDP‐choline (500 mg/kg; i.p.) for 14 days delayed cell
membrane damage and behavioral dysfunction in spontaneously hypertensive rats in which ischemia had
been caused by artiﬁcially induced occlusion of the lateral middle cerebral artery (Aronowski et al., 1996).
In a similar study, CDP‐choline (500 mg/kg; i.p.) decreased infarct volume and edema in a rat model of
temporary focal ischemia (Schabitz et al., 1996). In mice with an intracerebral hemorrhage, CDP‐choline
(500 mg/kg; i.p.) reduced the volume of ischemic injury surrounding the hematoma, and improved the
behavioral outcome (Clark et al., 1998). In another study, CDP‐choline (400 mg/kg; i.p.) increased blood
pressure, reduced infarct volume, and decreased the mortality rates of hypotensive rats with a experimental
subarachnoid hemorrhage (Alkan et al., 2001). In rats with permanent occlusion of the middle cerebral
artery, CDP‐choline inhibited MAP kinase signaling pathways (Krupinski et al., 2005). In a focal brain
ischemia model in rats, CDP‐choline (0.5–2 g/kg; i.p.) reduced infarct size and inhibited ischemia‐induced
decreases in cortical and striatal ATP levels (Hurtado et al., 2005).
CDP‐choline produces synergistic neuroprotective effects when this treatment is combined with
glutamate receptor antagonists (i.e., MK‐801; Onal et al., 1997 or lamotrigine; Ataus et al., 2004);
thrombolytic agents (i.e., recombinant tPA; Andersen et al., 1999; De Lecinana et al., 2006 or urokinase;
Shuaib et al., 2000); the calcium channel blocker, nimodipine (Sobrado et al., 2003); or basic ﬁbroblast
growth factor (Schabitz et al., 1999) using experimental ischemia models in rats.
In a gerbil model in which brain ischemia was produced by bilateral ligation of the carotid arteries,
intraventricular (0.6 mmol; Trovarelli et al., 1981) or intraperitoneal (150 mg/kg; Trovarelli et al., 1982)
CDP‐choline partially prevented the ischemia‐induced increases in fatty acids and decreases in PC levels
(Trovarelli et al., 1981, 1982). CDP‐choline reduced the dysfunctions of the BBB after reperfusion in gerbils
(Rao et al., 1999), and reduced the cerebral edema, concurrently reducing the levations of AA levels and
36 3.2 Choline and its products acetylcholine and phosphatidylcholine
leukotriene C4 synthesis (Rao et al., 2000). In a transient cerebral ischemia model, CDP‐choline (500 mg/kg
daily for 2 days; i.p.) restored the decreases in PC, SM, cardiolipin, and total glutathione levels induced by
ischemia (Adibhatla et al., 2001).
In cats undergoing brief periods of cerebral ischemia, CDP‐choline attenuated the depression in the
cortical evoked potentials (Boismare et al., 1978).
7.2 Head Trauma (Cranio‐Cervical Trauma)
Neuroprotective effects of CDP‐choline have been described in studies using various trauma models and
experimental animals. In a weight‐drop concussive head injury model in mice, CDP‐choline (60–250
mg/kg) shortened the recovery time (Boismare et al., 1977). In a controlled lateral‐impact model in rat,
CDP‐choline (100 mg/kg; i.p.) increased extracellular ACh levels, decreased cognitive deﬁcits and attenu-
ated the trauma‐induced increased sensitivity to the memory‐disrupting effects of scopolamine (Dixon
et al., 1997). In a cortical impact model in rat, intraperitoneal CDP‐choline (400 mg/kg) reduced brain
edema (Baskaya et al., 2000), and decreased neuronal loss in the hippocampus, and improved neurological
recovery (Dempsey and Raghavendra Rao, 2003). In a rat experimental (weight‐drop) spinal cord injury
model, CDP‐choline (400 mg/kg; i.p.) improved behavioral and neuroanatomic signs of recovery (Yucel
et al., 2006).
7.3 Induced Lesions
Neuroprotective actions of CDP‐choline have also been demonstrated in lesion studies. Oral administration
of CDP‐choline, at a daily dose of 1 g/kg for 4 days, signiﬁcantly extended survival time and increased the
percentage of survivors from KCN‐induced toxicity (Tornos et al., 1983a). CDP‐choline administration
(500 mg/kg, i.p.) for 7 days ameliorated functional behavior, as shown by reducing the number of
apomorphine‐induced contralateral rotations. It also attenuated the loss of substantia nigra dopaminergic
neurons and the decrease in tyrosine hydroxylase immunoreactivity, in the ipsilateral striatum in rats
injected intrastriatally with the dopaminergic toxin, 6‐hydroxydopamine (Barrachina et al., 2003). CDP‐
choline (62.5–250 mg/kg, i.p.) protected hippocampal neurons against apoptosis and the degeneration
induced by injecting beta‐amyloid into brains of rats also undergoing cerebral hypoperfusion (Alvarez et al.,
1999). CDP‐choline (50 mg/kg) prevented mice and rats from an acrylamide‐induced neurological syn-
drome (Agut et al., 1983). In tissue culture studies, CDP‐choline protected the retinal ganglion cells
(Oshitari et al., 2002) and prevented glutamate‐mediated cell death in cerebellar granule neurons (Mir
et al., 2003).
7.4 Other Effects
Oral or intraperitoneal administration of CDP‐choline (10–500 mg/kg, for 5–7 days) improved memory in
rats with memory deﬁcits induced by muscarinic AChR antagonists, by the a2‐adrenoceptor agonist
clonidine, by electroconvulsive shock, or by hypoxia (Petkov et al., 1992, 1993). Dietary CDP‐choline
supplementation protected rats against the development of memory deﬁcits in aging (Teather and
Wurtman, 2003), and prevented memory impairments caused by impoverished environmental conditions
(Teather and Wurtman, 2005). In humans, CDP‐choline improved verbal memory in aging (Spiers et al.,
1996) and beneﬁtted memory in elderly subjects (Alvarez et al., 1997).
In rat striatum, CDP‐choline activated tyrosine hydroxylase (Martinet et al., 1981), increased dopamine
levels (Martinet et al., 1979; Shibuya et al., 1981), and enhanced K
‐evoked dopamine release (Agut et al.,
2000) and haloperidol‐induced elevation in dopamine metobolites (Agut et al., 1984). Oral CDP‐choline
increased the total urinary excretion of the noradrenaline metabolite 3‐methoxy‐4‐hydroxyphenylglycol, in
rats and humans (Lopez G‐Coviella et al., 1986). Centrally administered CDP‐choline (0.5–2.0 mmol)
Choline and its products acetylcholine and phosphatidylcholine 3.2 37
increased plasma vasopressin (Cavun et al., 2004), and ACTH concentrations and potentiated the release of
GH, TSH, and LH (Cavun and Savci, 2004) stimulated by clonidine, TRH, and LHRH, respectively.
Intravenously injected CDP‐choline (250 mg/kg) increased plasma concentrations of noradrenaline and
adrenaline in rats (Savci et al., 2003).
Oral administration of a single dose (2 g/kg) of CDP‐choline to mice decreased the intensity of the
morphine withdrawal syndrome (Tornos et al., 1983b). In a single study, treatment with CDP‐choline was
reported to affect some measures of craving in cocaine‐dependent human subjects (Renshaw et al., 1999).
CDP‐Choline increased blood pressure, reversed hypotension in hemorrhagic shock (Savci et al., 2002a,
2003), and prolonged survival time (Yilmaz et al., 2006b) when given intravenously (100–500 mg/kg) or
intracerebroventricularly (0.1–2.0 mmol) to rats.
CDP‐choline decreased platelet reactivity to aggregating agents when given acutely (250 mg/kg) and
increased the antiaggregatory activity of aortic walls when given chronically (250 mg/kg, i.p., 2 weeks) to
rats. In dogs, CDP‐choline (70 mg/kg; i.v.) prevented the endotoxin‐induced decrease in circulating platelet
counts and prolonged platelet closure times (Yilmaz et al., 2006a).
Intracerebroventicular CDP‐choline (0.5–2.0 mmol) produced antinociception in three different acute
pain models (i.e., thermal paw withdrawal tests, mechanical paw pressure test, and acetic acid writhing test)
in rats (Hamurtekin and Gurun, 2006).
7.5 Clinical Studies
CDP‐choline effects have been examined in studies involving numerous normal subjects and patients with
cerebral ischemia, traumatic brain injury, hypoxia, Alzheimer’s and Parkinson’s diseases. To date sufﬁcient
evidence has not been accumulated regarding any such use to warrant its approval for drug status by the US
Food and Drug Administration. It is, however, approved for sale in a few other countries and sold under its
international nonproprietary name, citicoline.
Clinical trials conducted in the USA, tested daily oral doses of 500, 1000, or 2000 mg/day, given for
6 weeks. In some such studies, the drug was administered within the ﬁrst 48 h of an ischemic stroke (Clark
et al., 1999); in others, it was ﬁrst given to patients up to 14 days after the onset of the ischemic episode
(Tazaki et al., 1988). Pooling of individual patients data from four USA trials yielded evidence that CDP‐
choline treatment could improve overall recovery at 12 weeks in acute ischemic stroke patients (Davalos
et al., 2002). Pooled diffusion‐weighted magnetic resonance imaging data from two clinical trials showed a
signiﬁcant dose‐dependent reduction on percent change in lesion volume (Warach, 2002).
8 Choline in Autonomic and Motor Neurons
All nerves that leave the brain or spinal cord (i.e., axons of motor neurons, parasympathetic preganglionic
neurons, and sympathetic preganglionic neurons), as well as all postganglionic parasympathetic neurons,
release ACh as their neurotransmitter, and in all of them choline availability determines the rates at which
ACh is synthesized and released. ACh is also present in the periphery in placenta, lymphocytes, the bladder,
and tracheal epithelium; however, in these cells the effects of increasing choline availability on ACh
synthesis have not yet been determined.
More than 50 years ago, Hutter (1952) demonstrated that a low intravenous dose (7 mg/kg) of choline
enhanced neuromuscular transmission, whereas a high dose (50 mg/kg) blocked this transmission. Hutter
also demonstrated that choline, at doses of 3–60 mg/kg, could restore neuromuscular transmission in
curarized cats. Based on these observations, he suggested that choline increased ACh output from motor
nerve endings. More recently, using the isolated, vascularly perfused rat phrenic nerve‐hemidiaphragm
preparation Bierkamper and Goldberg (1979, 1980) directly demonstrated that choline (at 30–60 mM
concentrations) could increase ACh release at the neuromuscular junction.
Effects of choline on ACh synthesis and release, and on cholinergic neurotransmission, at para-
sympathetic synapses have been demonstrated in vivo (Kuntscherova, 1972; Haubrich et al., 1974, 1975;
38 3.2 Choline and its products acetylcholine and phosphatidylcholine
Ilcol et al., 2003a) and in vitro, using isolated hearts (Dieterich et al., 1978), and atrial (Meyer and Baker,
1986) and pancreatic minces (Ilcol et al., 2003a). Choline infusion (10 mM) increases by 2–3‐fold ACh
release evoked by electrical stimulation of the vagus nerve in chicken hearts, and by at least 23‐fold in cat
heart (Dieterich et al., 1978). The presence of choline (10 mM) in the perfusion medium also increased,
by—two to threefold, ACh release evoked by electrical ﬁeld stimulation (at 20 Hz for 20 min) from isolated
chicken, rat, cat, and guinea pig hearts (Dieterich et al., 1978). Subcutaneous choline administration (200
mg/kg) increased, by 34%, the ACh content of the atrium (Kuntscherova, 1972) and in atrial minces,
choline (at 1–100 mM) increased ACh synthesis and release in a concentration‐dependent manner (Meyer
and Baker, 1986). In rats, intraperitoneal choline (90 mg/kg) increased, by 45%, the ACh contents of
pancreatic tissue; this was associated with increased cholinergic neurotransmission to insulin secreting
b‐cells (Ilcol et al., 2003a). Choline (10–130 mM) also increased ACh synthesis and release from rat
pancreatic minces (Ilcol et al., 2003a).
Oral choline administration (20 mmol/kg) to rats increases adrenal ACh levels by more than twofold for
8 h (Ulus et al., 1977a) and tyrosine hydroxylase activity by about 30% (Ulus et al., 1977a). Repeated oral
administration of choline for 4 days increases the enzyme activity further, by up to 50–60%; the increase in
tyrosine hydroxylase activity is not observed after intubation with saline, water, or ammonium chloride,
and fails to occur in rats pretreated with cycloheximide (Ulus et al., 1977a). Similar increases in tyrosine
hydroxylase activity after oral choline administration are observed in sympathetic ganglia, including the
superior cervical ganglion, the stellate and celiac ganglia (Ulus et al., 1977c, 1979; Ulus and Wurtman,
1979), and the ganglia of the thoracic sympathetic chain (Ulus et al., 1977c). These increases in tyrosine
hydroxylase activity are also not seen in adrenals after adrenal denervation (Ulus et al., 1977a) or after
decentralization of superior cervical ganglion (Ulus et al., 1979), indicating that the action of choline is
transsynaptic and that it requires intact preganglionic cholinergic nerves to affect ACh synthesis and release.
Further evidence that choline administration enhances ACh release was obtained by studies in which
choline was administered along with reserpine, or with other drugs that increase impulse ﬂow in pregan-
glionic cholinergic nerves. Injection of reserpine (2.5 mg/kg; i.p.) phenoxybenzamine (20 mg/kg; i.p.), or
insulin (2 units/rat; i.p.) daily for 4 days, or of 6‐hydroxydopamine (200 mg/kg twice, with an interval of
48 h, through tail vein), all caused marked increases in adrenal tyrosine hydroxylase activities. When these
treatments were combined with oral choline (2.8 g/kg; by stomach tube), the resulting increases in tyrosine
hydroxylase activity were considerably greater than the sum of the changes produced by choline alone and
drug alone, that is, signiﬁcant potentiation occurred (Ulus et al., 1977a, b, 1978). Potentiation of a
treatment‐induced rise in tyrosine hydroxylase by choline was also observed in the adrenals of rats kept
in the cold (Ulus et al., 1977a, 1978) and in superior cervical ganglia of reserpine‐treated rats (Ulus et al.,
1977a). Taken together, these data indicate that the availability of free choline is a major factor controlling
cholinergic neurotransmission in the sympathoadrenal system.
Studies using the classical perfused‐superior cervical ganglion system failed to demonstrate parallel
increases in ACh release in response to elevating the choline concentration of the perfusion media (Birks
and MacIntosh, 1961; Matthews, 1966; MacIntosh, 1979; O’Regan and Collier, 1981). However, when
superior cervical ganglia were perfused with choline‐containing (10–14 mM) plasma or with Locke solution,
they released greater amounts of ACh, by twofold, during a 1‐h stimulation (20 Hz) period than ganglia
superfused without exogenous choline (Birks and MacIntosh, 1961; Matthews, 1966; MacIntosh, 1979).
Furthermore, the ACh stores in perfused‐ganglia stimulated without exogenous choline were found to be
partly depleted, although these ganglia had managed to synthesize some ACh by reusing free choline
generated from the hydrolysis of released ACh, or from hydrolyzing membrane phospholipids like PC). PC
levels and the number of synaptic vesicles in the cat’s superior cervical ganglion were found to decline
signiﬁcantly after stimulation of the preganglionic nerve trunk if the uptake of exogenous choline was
blocked by HC‐3 (Parducz et al., 1976), or if the ganglia were perfused with a choline‐free Locke solution
(Parducz et al., 1986). In striking contrast, stimulated ganglia supplied with exogenous choline maintained
their ACh stores (Birks and MacIntosh, 1961; Matthews, 1966; MacIntosh, 1979), as well as membrane PC
levels and the numbers of storage vesicles (Parducz et al., 1976, 1986), although they released much more
ACh than they initially contained.
Choline and its products acetylcholine and phosphatidylcholine 3.2 39
The authors thank Dr. Jan Krzysztof Blustajn and Ms. Carol Watkins for the critical review of this chapter.
Studies described in this chapter were supported in part by grants from the National Institutes of Mental
Health (MH‐28783); the NIH‐NCRR (5‐MO1RR01066–29); the Center for Brain Sciences and Metabolism
Charitable Trust; and the Turkish Academy of Sciences (Ismail H. Ulus).
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