Revealing CD38 cellular localization using a cell permeable, mechanism-based fluorescent small-molecule probe

Article abstract of DOI:10.1021/ja411046j

Nicotinamide adenine dinucleotide (NAD) is increasingly recognized as an important signaling molecule that affects numerous biological pathways. Thus, enzymes that metabolize NAD can have important biological functions. One NAD-metabolizing enzyme in mammals is CD38, a type II transmembrane protein that converts NAD primarily to adenosine diphosphate ribose (ADPR) and a small amount of cyclic adenosine diphosphate ribose (cADPR). Localization of CD38 was originally thought to be only on the plasma membrane, but later reports showed either significant or solely, intracellular CD38. With the efficient NAD-hydrolysis activity, the intracellular CD38 may lead to depletion of cellular NAD, thus producing harmful effects. Therefore, the intracellular localization of CD38 needs to be carefully validated. Here, we report the synthesis and application of a cell permeable, fluorescent small molecule (SR101-F-araNMN) that can covalently label enzymatically active CD38 with minimal perturbation of live cells. Using this fluorescent probe, we revealed that CD38 is predominately on the plasma membrane of Raji and retinoic acid (RA)-treated HL-60 cells. Additionally, the probe revealed no CD38 expression in K562 cells, which was previously reported to have solely intracellular CD38. The finding that very little intracellular CD38 exists in these cell lines suggests that the major enzymatic function of CD38 is to hydrolyze extracellular rather than intracellular NAD. The fluorescent activity-based probes that we developed allow the localization of CD38 in different cells to be determined, thus enabling a better understanding of the physiological function.

Full text of DOI:10.1021/ja411046j

Article
pubs.acs.org/JACS
Revealing CD38 Cellular Localization Using a Cell Permeable,
Mechanism-Based Fluorescent Small-Molecule Probe
Jonathan H. Shrimp,† Jing Hu,† Min Dong,† Brian S. Wang,† Robert MacDonald,‡ Hong Jiang,†
Quan Hao,§,‡ Andrew Yen,‡ and Hening Lin*,†
†Department of Chemistry and Chemical Biology, Cornell University, Ithaca, New York 14853, United States
‡Department of Biomedical Sciences, Cornell University, Ithaca, New York 14853, United States
§Department of Physiology, University of Hong Kong, Hong Kong, China
S
* Supporting Information
ABSTRACT: Nicotinamide adenine dinucleotide (NAD) is
increasingly recognized as an important signaling molecule that
affects numerous biological pathways. Thus, enzymes that
metabolize NAD can have important biological functions. One
NAD-metabolizing enzyme in mammals is CD38, a type II
transmembrane protein that converts NAD primarily to adenosine
diphosphate ribose (ADPR) and a small amount of cyclic
adenosine diphosphate ribose (cADPR). Localization of CD38
was originally thought to be only on the plasma membrane, but
later reports showed either significant or solely, intracellular CD38.
With the efficient NAD-hydrolysis activity, the intracellular CD38
may lead to depletion of cellular NAD, thus producing harmful
effects. Therefore, the intracellular localization of CD38 needs to
be carefully validated. Here, we report the synthesis and application of a cell permeable, fluorescent small molecule (SR101−F-
araNMN) that can covalently label enzymatically active CD38 with minimal perturbation of live cells. Using this fluorescent
probe, we revealed that CD38 is predominately on the plasma membrane of Raji and retinoic acid (RA)-treated HL-60 cells.
Additionally, the probe revealed no CD38 expression in K562 cells, which was previously reported to have solely intracellular
CD38. The finding that very little intracellular CD38 exists in these cell lines suggests that the major enzymatic function of CD38
is to hydrolyze extracellular rather than intracellular NAD. The fluorescent activity-based probes that we developed allow the
localization of CD38 in different cells to be determined, thus enabling a better understanding of the physiological function.
However, CD38 catalyzes the formation of mainly adenosine
diphosphate ribose (ADPR) and only a minute amount of
cADPR.7−10 Under certain conditions, CD38 can also catalyze
a base-exchange reaction converting nicotinamide adenine
dinucleotide phosphate (NADP) to nicotinic acid adenine
dinucleotide phosphate (NAADP), which is also a calcium
mobilizing messenger.11 As a receptor, it was reported that
CD38 can initiate transmembrane signaling in response to
antibody binding.12
The cellular localization of CD38 has also been perplexing.
CD38 was originally identified as a cell surface protein, but was
later reported to be present in intracellular compartments, such
as the mitochondria, Golgi, and ER, with the highest levels of
expression in the nuclear membranes.13−20 In addition, a recent
report suggested the existence of type III CD38 on the plasma
membrane with the catalytic domain facing the cytosol.21
However, conflicting results on CD38 cellular localization have
been reported, with some showing only plasma membrane
INTRODUCTION
■
Nicotinamide adenine dinucleotide (NAD) is an important
cofactor used in many metabolic reactions. Recently, it has been
increasingly recognized that, in addition to serving as a cofactor,
it also serves as an important signaling molecule by effecting
protein posttranslational modifications, such as NAD-depend-
ent deacylation and ADP-ribosylation.1,2 Thus, enzymes that
metabolize NAD can have important biological functions. One
mammalian enzyme that metabolizes NAD is cluster of
differentiation 38 (CD38). CD38, a type II membrane protein,
has important physiological functions, demonstrated by the
compromised immune response and social memory defect in
CD38 knockout mice.3,4 In addition, its expression is associated
with a poor prognosis in chronic lymphocytic leukemia.5
However, the molecular mechanism underlying its physiological
functions is still not well understood. It is reported to function
as both an enzyme and a receptor. CD38 has 69% overall
homology to an Aplysia cyclase, which converts NAD to cyclic
adenosine diphosphate-ribose (cADPR), a molecule that is
reported to be a calcium mobilizing messenger.6 CD38 was
thus considered to have a similar enzymatic function.6
Received: October 29, 2013
© XXXX American Chemical Society
A
dx.doi.org/10.1021/ja411046j | J. Am. Chem. Soc. XXXX, XXX, XXX−XXX

Journal of the American Chemical Society
Article
CD38 and others showing only nuclear localized CD38.15,22
Because of the efficient NAD-hydrolysis activity of CD38, the
intracellular CD38 may lead to depletion of cellular NAD, thus
producing detrimental effects. Therefore, the intracellular
localization of CD38 merits careful investigation. Methods
used to study cellular localization of CD38 include the use of
antibodies for confocal immunofluorescence after cell fixation
and permeabilization, subcellular fractionation, and less
frequently, CD38−GFP fusion proteins.13−20,22 To mitigate
the possibility of artifacts from these methods, there is a need
for new methods with minimal perturbation of live cells and
minimal interference with CD38 signaling. In addition,
previously used labeling methods are typically followed by a
separate step of organelle isolation to check CD38 enzymatic
activity. A method to both localize and demonstrate activity will
be helpful.
fluorescent activity-based small-molecule probe for labeling
CD38 in live cells. This probe allowed detection of catalytically
active CD38 both on the plasma membrane and inside the cells.
Using this probe, we investigated the cellular distribution of
CD38 in two leukemia (HL-60 and K562) and one lymphoma
(Raji) cell lines that were previously reported to have
intracellular CD38.15,16 Our results showed CD38 is localized
mainly at the plasma membrane with very little intracellular
CD38. In particular, our results showed no pronounced nuclear
localization as previously reported, which may have been an
artifact of the immunofluorescence method. The clarification of
CD38 intracellular localization will facilitate the understanding
of its physiological function.
EXPERIMENTAL SECTION
■
General Methods. Reagents, unless specified otherwise, were
purchased from commercial suppliers in the highest purity available
and used as supplied. Antibodies for detection of human CD38
(mouse) were bought from BD Biosciences (San Jose, California,
USA, cat. #: 611114), while antibodies for detection of human
GAPDH (rabbit), as well as horseradish peroxidase-linked antimouse
and antirabbit antibodies were bought from Cell Signaling (Danvers,
MA, USA). Mammalian protein extraction reagent (M-PER) was
bought from Pierce (Rockford, IL, USA). All-trans retinoic acid (RA),
protease and phosphatase inhibitors were bought from Sigma (St.
Louis, MO, USA). ECL was bought from GE Healthcare (Pittsburgh,
PA, USA). Both sulforhodamine and tetramethylrhodamine fluo-
rescence signals from protein gels were recorded by Typhoon 9400
Variable Mode Imager (GE Healthcare Life Sciences). For visual-
ization of tetramethylrhodamine, a green laser at 532 nm was used for
excitation and a collection filter of 580 nm with band-pass of 30 nm
was used for emission collection. For visualization of sulforhodamine,
the 532 nm laser was used for excitation and 610BP30 filter for
emission collection. Detection of fluorescence was done using a PMT
at 650 V and normal sensitivity. Images collected were analyzed by
ImageQuant TL v2005.
Previously, we developed a CD38-labeling method that uses
a suicide substrate, 2′-deoxy-2′-fluoro arabinosyl NAD
conjugated with tetramethylrhodamine, Rh-6-(F-araNAD)
(Scheme 1A). Rh-6-(F-araNAD) can covalently label CD38
Scheme 1. (A) Molecular Structure of Rh-6-(F-araNAD);
(B) Synthesis of the Cell-Permeable Fluorescent CD38
Probe, SR101−F-araNMN
Labeling of Purified CD38 (wt and E226D, E226Q mutants)
with SR101−F-araNMN In Vitro. CD38 (wt or E226D and E226Q
mutants, 8 μM, expressed and purified as reported earlier24,25) and
sulforhodamine 101-F-araNMN (SR101−F-araNMN) (20 μM) in 10
μL reaction buffer (25 mM HEPES, 50 mM NaCl, pH 7.4) were
incubated at 37 °C for 30 min, then mixed with 2 μL 6× protein
loading buffer. The samples were heated at 100 °C for 7 min and then
resolved by SDS-PAGE. Before staining with Coomassie blue, the gel
was irradiated under UV light (Transillum, Fisher Scientific, model
DLT-A), and the fluorescence image was recorded with a digital
camera (Nikon Coolpix L22).
Labeling of CD38 in Live Cells. HL-60 cells were treated with 1
μM RA in cell culture media (GIBCO RPMI Medium 1640 with 10%
GIBCO Heat-inactivated Fetal Bovine Serum) for 24 h in a 5% CO2
incubator at 37 °C. Untreated HL-60, Raji, and K562 cells were
cultured using the same media without RA. Then the cells were
harvested from 4 mL cell culture (1 × 106 cells/mL) by centrifugation
at 25 °C, 1200 rpm for 5 min. The cells were initially washed once
using 500 μL PBS. Cells were resuspended in 100 μL PBS (reaction
volume). In one experimental procedure, these cells were directly
labeled with SR101−F-araNMN as detailed below to label all the
CD38 molecules. In another experimental procedure, the cells were
first treated with a nonfluorescent, cell-impermeable probe, 6-alkyne-F-
araNAD, to block the cell surface CD38 before labeling intracellular
CD38 with SR101−F-araNMN. 6-Alkyne-F-araNAD was added to a
final concentration of 10 μM. After incubation at RT for 8 min, the
cells were washed once with 500 μL PBS. Cells were resuspended in
100 μL PBS (reaction volume), and SR101−F-araNMN was added to
a final concentration of 10 μM and allowed to incubate at RT for 8
min. The cells were then washed once with 500 μL cold PBS (PBS at 4
°C), followed by resuspending the cells in 1 mL cold methanol
(methanol stored in −20 °C for at least 1 h prior to use), and samples
on the key catalytic residue, Glu226, forming a stable covalent
intermediate.23 This method was successful in labeling CD38
while not interfering with antibody-induced CD38 signaling
events. It was mechanism-based labeling and thus only labeled
catalytically active CD38. However, this molecule was not cell
permeable; therefore, it cannot be used to label intracellular
CD38. In the present study, we developed a cell permeable
B
dx.doi.org/10.1021/ja411046j | J. Am. Chem. Soc. XXXX, XXX, XXX−XXX

Journal of the American Chemical Society
Article
were held at −20 °C for 10 min. Methanol was removed after
centrifugation at 4 °C, 2000 rpm to pellet the cells. Cells were then
resuspended in a fresh 1 mL of cold methanol (methanol at −20 °C)
and incubated on ice for 40 min. Again, methanol was removed and
cells were washed once with 500 μL fresh, cold PBS (PBS at 4 °C) to
ensure removal of methanol. Finally, cells were resuspended in 100 μL
of PBS. Then, 10 μL PBS containing the cell suspension was applied
onto a microscope slide and covered with a micro cover glass.
Confocal images (8 line average) of cells were acquired with a Zeiss
LSM 710 confocal microscope with a 63×/1.4 oil immersion objective.
Green 561 nm (15mW DPSS laser, laser power percentage given in
the figure captions) was used for sulforhodamine (SR) fluorescence.
Emission signal in the range of 566−717 nm (SR emission) was
detected.
dry nonfat milk in PBS-Tween before probing with 1:1000 (v/v)
dilutions of antibody in 5% BSA in PBS-Tween overnight at 4 °C.
Membranes were probed with 1:1000 (v/v) dilutions of secondary
antibody in 5% BSA in PBS-Tween for 1 h at RT before visualizing
with ECL. Blots shown are representative of at least three independent
repeats.
RESULTS
■
Design and Synthesis of a Cell-Permeable CD38
Probe. Our strategy for making a cell-permeable CD38
probe was to first choose a cell-permeable fluorophore and
then conjugate it to F-araNMN instead of F-araNAD (Scheme
1). F-araNMN is smaller and has one fewer negative charges
than F-araNAD, and thus may be more cell permeable. Among
several fluorescent dyes that we checked, sulfurhodamine 101
(SR101) showed the desired cell permeability. This was
interesting as SR101 has size, charge, and overall structure
similar to those of tetramethylrhodamine, yet SR101 is more
cell permeable. To conjugate SR101 to F-araNMN, we decided
to use the copper-catalyzed Huisgen 1,3-dipolar cycloaddition
between alkyne and azide, commonly known as click chemistry
(Scheme 1B).26 We designed an alkyne-containing F-araNMN
compound (F-araNMN alkyne) and SR101 azide compound.
To obtain F-araNMN alkyne, we began with a 2′-fluoroarabino-
side (1) where all hydroxyl groups were protected with benzoyl
groups. The anomeric O-benzoyl group was first replaced with
bromine and then by nicotinamide. The benzoyl groups were
removed using potassium carbonate in methanol. Then in a
one-pot reaction, the 5′-hydroxyl group was phosphorylated
and then connected to 3-butyn-1-ol via a phosphodiester bond
to give the desired F-araNMN alkyne compound. To make the
SR101 azide compound, we first made the sulfonyl chloride
derivative of SR101 and then attached a linker with an amino
group and an azido group at opposite ends. Finally, the SR101
azide was conjugated to F-araNMN alkyne via click chemistry
to obtain the desired SR101−F-araNMN (Scheme 1B). The
detailed synthesis is shown in Scheme S1 in the Supporting
Information (SI).
In-Gel Fluorescence Analysis of CD38 Intracellular Local-
ization. Lysis buffer recipe: Tris-HCl pH 7.9 (25 mM), NaCl (150
mM), glycerol (10%), Igepal (1%), 25 μL protease inhibitor cocktail
(PIC, Sigma-Aldrich, #P8340)/500 uL, PMSF (0.5 mM), EDTA pH
8.0 (5 mM). HL-60 cells were treated with 1 μM RA in cell culture
media for 24 h in a 5% CO2 incubator at 37 °C. Raji and K562 cells
were not treated with RA. Untreated HL-60, Raji, and K562 cells were
cultured using the same media except for being used without any
added treatment. Similar amounts of cells were used for both live cell
and whole cell lysate in-gel fluorescent labeling. To do this, cells were
counted to make sure the same number of cells were used in the live
cell and the whole cell lysate labeling. Cells were harvested by
centrifugation at 25 °C, 1200 rpm, for 5 min. Cells were washed twice
with 1 mL PBS. From this point, one batch of cells was lysed to obtain
whole cell lysate, and then the fluorescent labeling was carried out, and
another batch of cells was fluorescently labeled first, and then whole
cell lysate was collected. To label live cells, the cells were first
suspended in 100 μL PBS (reaction volume) and fluorescent
moleculeeither SR101−F-araNMN or Rh-6-(F-araNAD)was
added to a final concentration of 10 μM. Then the sample was kept
at RT for about 8 min, followed by centrifugation at 4 °C, 1500 rpm,
for 3 min. Cells were washed twice using 500 μL cold PBS (PBS at 4
°C). PBS was removed prior to addition of about 30 μL lysis buffer
followed by freeze/thaw lysis (the samples were frozen at −80 °C and
then removed from −80 °C and thawed on ice for ∼30 min while
vortexing briefly every 5 min). Once samples were fully thawed on ice,
they were centrifuged at 4 °C, 14,000 rpm, for 6 min to collect the
supernatant (proteins solubilized by detergent in lysis buffer). Protein
concentration in the cell lysate was determined using the Bradford
assay, and 25 μg of lysate was resolved by SDS-PAGE. In the case of
collecting whole cell lysate followed by fluorescent labeling, the cells
were washed with PBS and resuspended in 30 μL lysis buffer, and then
the whole cell lysates were collected as described above. Protein
concentration in the cell lysate was determined using the Bradford
assay. Once protein concentration was determined, stock lysate was
diluted to 2.5 μg/μL, and 10 μL of the diluted lysates was mixed with
the fluorescent molecule to give a final 10 μM labeling concentration.
The fluorescent labeling reaction was quenched using 2 μL of 6× SDS
containing protein loading buffer. The samples were heated at 100 °C
for 7 min and then resolved by SDS-PAGE. Before staining with
Coomassie blue, the fluorescence image of the gel was recorded by a
Typhoon 9400 Variable Mode Imager with settings of green laser at
532 nm and emission collection filter of 580BP30 (rhodamine) and
610BP30 (sulforhodamine). Detection was done using PMT650 V
(normal sensitivity), and data were analyzed by ImageQuant TL
v2005.
In Vitro Labeling of Purified CD38 by SR101−F-
araNMN. To test whether SR101−F-araNMN can covalently
label CD38 at the catalytic E226 residue, we used purified, wild-
type, and catalytic mutants (E226Q and E226D) of CD38
extracellular catalytic domain. Both wild-type and mutants were
incubated with SR101−F-araNMN for 10 min. The reaction
mixtures were resolved by SDS-PAGE and visualized by
fluorescence and then stained with Coomassie blue. Wild-
type CD38 was fluorescently labeled by SR101−F-araNMN,
but the CD38 catalytic mutants were not (Figure 1). These
results demonstrated that SR101−F-araNMN is an activity-/
mechanism-based probe for CD38. We compared the labeling
efficiency of Rh-6-(F-araNAD) and SR101−F-araNMN. With
CD38 at 1 μM and the probes at 10 μM, the labeling reactions
were complete within 0.5 min for Rh-6-(F-araNAD) and within
Western Blot Analysis. Cell cultures were seeded at 0.2 × 106
cells/mL, and those that received RA were treated at a concentration
of 1 μM. After 48 h, cells were washed twice with PBS before being
lysed by adding 400 μL of a lysis buffer consisting of M-PER with
1:100 (v/v) dilutions of protease inhibitors and phosphatase
inhibitors, and the cells were placed on ice for 30 min. The lysate
was then spun at 4 °C, 13,000 rpm, for 30 min, and supernatant was
saved and used for analysis. Then, 25 μg of protein lysate was loaded
per lane and resolved by SDS-PAGE analysis and transferred to PVDF
membrane. Membranes were then blocked for 1 h in a solution of 5%
Figure 1. In-vitro labeling of purified CD38 with SR101−F-araNMN.
Lanes 1 and 2, CD38 wt; lanes 3 and 4, CD38 E226D; lanes 5 and 6,
CD38 E226Q. Ladder is on the left, listed first.
C
dx.doi.org/10.1021/ja411046j | J. Am. Chem. Soc. XXXX, XXX, XXX−XXX

Journal of the American Chemical Society
Article
5 min for SR101−F-araNMN (Figure S1 in SI). Furthermore,
the addition of NAD (1 mM) or nicotinamide (100 μM) to the
labeling reaction had very little effect on labeling efficiency
(Figure S1). Thus, SR101−F-araNMN is slightly less efficient
than Rh−F-araNAD, but still labels CD38 very efficiently.
To confirm that this probe is specific for CD38 in cells, we
used the human leukemia cell line, HL-60. HL-60 was chosen
because these cells have a very low levels of CD38 but could be
induced to express higher levels of CD38 with RA.27
Consequently, untreated HL-60 cells were used as the negative
control. In one experiment, we labeled live cells, followed by
collecting whole cell lysates and then resolved the whole cell
lysates with SDS-PAGE (lanes 1, Figure 2). In another
Figure 2. In-gel fluorescence analysis of CD38 in HL-60 cells (RA and
untreated). Lanes 1, live-cell labeling with SR101−F-araNMN; lanes 2,
whole cell lysate labeled with SR101−F-araNMN; lanes 3, whole cell
lysate with no CD38 probe. Ladder is on the left, listed first.
Figure 3. Confocal images of HL-60 cells (with or without RA-
treatment): (A) Cells without RA-treatment labeled with SR101−F-
araNMN and washed with PBS. (B) Cells without RA-treatment
labeled with Rh-6-(F-araNAD) and washed with PBS. (C) RA-treated
cells labeled with SR101 (negative control) and washed with
methanol. (D) RA-treated cells labeled with SR101−F-araNMN and
washed with methanol. Confocal microscope settings: laser power:
4.5%, pinhole: 1.1 airy unit, master gain for PMT: 875.
experiment, we collected the whole cell lysate, incubated with
the probe, and then resolved the lysates by SDS-PAGE (lanes 2,
Figure 2). In both cases, only one major fluorescent band,
which corresponds to the size of CD38, was detected in RA-
treated cells; while no major fluorescent bands were detected in
untreated cells (Figure 2). The weak fluorescent band around
20 kD was shown to be from a cleaved form of CD38 (Figure
S2 in SI). A weak fluorescent band at about 70 kD was also
detected in both RA treated and untreated HL-60 cells in the
live-cell labeling but not in the whole-cell lysate labeling (Figure
2). By optimizing the experimental conditions, we were able to
partially or completely eliminate this band (Figure S2 and S6 in
SI). Altogether, this result demonstrated that SR101−F-
araNMN is specific for CD38 in cells. The higher CD38
expression in RA-treated cells was also confirmed by Western
blot using a monoclonal anti-CD38 antibody (see Figure 7c).
Confocal Microscopy Imaging for Labeling of CD38 in
Live Cells with SR101−F-araNMN. The ability of SR101−F-
araNMN to enter live cells and label CD38 was next tested
using HL-60 cells and visualized using confocal microscopy. We
first incubated cells with either 10 μM SR101−F-araNMN or
10 μM Rh-6-(F-araNAD) for 8 min at RT followed by washing
with PBS to remove excess unbound dye. The incubation time
was determined to be sufficient to allow for the binding of the
probe to CD38 as determined by an in vitro labeling experiment
(Figure S1 in SI). We indeed found that SR101−F-araNMN
was cell permeable as strong fluorescence was observed inside
HL60 cells (Figure 3A); conversely, HL-60 cells labeled with
Rh-6-(F-araNAD) showed no fluorescence inside (Figure 3B).
However, washing with PBS alone could not remove excess
SR101−F-araNMN molecules in the cells (even HL-60 cells
without RA treatment had strong fluorescence inside), making
it difficult to differentiate free vs CD38-bound SR101−F-
araNMN. To circumvent this problem, we used methanol to
wash away the unbound SR101−F-araNMN as it was more
soluble in methanol. This was proven successful as HL-60 cells
treated with fluorescent molecule alone, SR101, then washed
with methanol showed essentially no fluorescence (Figure 3C).
In contrast, RA-treated HL-60 cells were strongly fluorescently
labeled with SR101−F-araNMN (Figure 3D) on the plasma
membrane and only weakly inside the cells.
Then, we turned our focus to locating the intracellular CD38
with SR101−F-araNMN. Although there was some weak
fluorescence inside the RA-treated HL-60 cells, it was difficult
to visualize the intracellular signal because the fluorescence on
the plasma membrane was too strong. To overcome this, we
used 6-alkyne-F-araNAD to first block plasma membrane
CD38. The 6-alkyne-F-araNAD was not cell permeable and had
no fluorescent molecule attached. HL-60 cells (RA treated and
untreated) were first incubated with 6-alkyne-F-araNAD to
block plasma membrane CD38, followed by incubation either
with SR101 as negative control or SR101−F-araNMN to label
intracellular CD38. This allowed us to clearly see the
intracellular fluorescently labeled CD38, after optimizing the
confocal microscope settings to maximize detection. HL-60
cells without RA treatment also showed very weak fluorescent
signal. This represented the low levels of CD38 present, which
was further confirmed by Western blot data shown later (see
D
dx.doi.org/10.1021/ja411046j | J. Am. Chem. Soc. XXXX, XXX, XXX−XXX

Journal of the American Chemical Society
Article
Figure 7C). Importantly, an increase in fluorescence (∼2.5-
fold) was observed in RA-treated cells compared to HL-60 cells
without RA treatment, confirming that the probe was labeling
CD38 (Figure 4).
after washing away excess probes (live cell labeling). The other
portion was lysed first and then the total cell lysate was
incubated with Rh-6-(F-araNAD) (whole cell lysate labeling).
The two batches of cell lysates were then resolved by SDS-
PAGE and analyzed by fluorescence and Coomassie blue
staining. If a significant amount of CD38 is intracellular, then
we expected that the live cell labeling intensity would be weaker
than the whole cell lysate labeling intensity with the
impermeable Rh-6-(F-araNAD). Contrary to this, we saw
essentially the same labeling intensities with the live cell
labeling and the whole cell lysate labeling (after correcting for
protein loading using the Coomassie blue staining) with Rh-6-
(F-araNAD) (Figure 5). In addition, to more accurately
Figure 4. Confocal images of HL-60 cells (RA or untreated) blocked
with 6-alkyne-(F-araNAD) followed by intracellular CD38 labeling
with SR101−F-araNMN. (A) RA treated HL-60 cells. (B) Untreated
HL-60 cells. Confocal microscope settings: laser power: 11.0%,
pinhole: 1.3 airy units, Master gain for PMT: 940. With laser settings
kept constant between the two images, higher fluorescence indicates
higher amount of active CD38.
Figure 5. In-gel fluorescence analysis showed that most CD38 was
present on plasma membrane. Lanes 1, live cell labeling with Rh-6-(F-
araNAD) followed by in-gel fluorescence (labeling plasma membrane
CD38 only); lanes 2, whole cell lysate was obtained first followed by
labeling with Rh-6-(F-araNAD) (labeling all catalytically active
CD38); lanes 3, whole cell lysate without CD38 probes. Protein
ladder is on the left, listed first. The full gel images are shown in Figure
S7 in SI.
It has been reported that cell fixation with methanol alone
can cause loss of cytosolic and nuclear proteins.28 To rule out
that the low detection of intracellular CD38 was not due to the
loss of CD38 during the methanol wash, we also used a
combination of paraformaldehyde (PFA) fixation with
methanol permeabilization after SR101−F-araNMN labeling
of CD38 in RA-treated HL-60 cells.28 This method preserves
both cell-surface and intracellular proteins.28 The data from this
combined PFA fixation and methanol permeabilization led to
the same conclusion that CD38 was mainly concentrated on
the plasma membrane (Figure S4 in SI). In addition, subcellular
fractionation of RA-treated HL-60 cells was done to obtain
nucleus, membrane, and cytosolic fractions.29,30 Using a
monoclonal anti-CD38 antibody that would presumably detect
both catalytically active and inactive CD38, the Western blot
data again showed that the nucleus fraction contained very low
levels of CD38, while the membrane fraction contain very high
levels of CD38 (Figure S5 in SI).
quantify the percentage of intracellular CD38, we used another
approach with the small-molecule probes. We first blocked cell
surface CD38 with 6-alkyne-F-araNAD and then labeled
intracellular CD38 with SR101−F-araNMN. In the control
experiment, we labeled the total CD38 using SR101−F-
araNMN without blocking of the cell surface CD38. We then
collected the whole cell lysates and used in-gel fluorescence to
quantify the intracellular CD38 vs total CD38 (Figure S3 in
SI). Quantification using this method showed that about 6.5%
of CD38 molecules were intracellular in RA-treated HL-60
cells. This result further supported the previous confocal
imaging results with SR101−F-araNMN and suggested that the
amount of intracellular and catalytically active CD38 was very
little.
CD38 Labeling in Raji and K562 Cells. Once cell
permeability and CD38 specific labeling was confirmed for
SR101−F-araNMN in HL-60 cells, we investigated the
intracellular distribution of CD38 in Raji and K562, which
are lymphoma and leukemia cell lines, respectively. Both types
of cells were reported to have intracellular CD38.15 In
particular, in Raji cells, it was reported that CD38 was present
in a special subnuclear location called the Cajal body. In K562
cells, it was reported that there was only intracellular CD38 and
no plasma membrane CD38.15
Thus, SR101−F-araNMN was cell permeable and capable of
labeling CD38 within live cells without perturbation of the cell
before labeling. In addition, it signifies that the intracellular
CD38 is in fact catalytically active. Despite the intracellular
presence, our results revealed that CD38 was mainly
concentrated on the plasma membrane as the plasma
membrane had the strongest fluorescence.
In-Gel Fluorescence of Whole Cell Lysate Using CD38
Probes to Label and Quantify Intracellular CD38. The
weak intracellular labeling diverged from previous reports
regarding pronounced intracellular CD38. We thus decided to
quantify the relative amount of intracellular vs plasma
membrane CD38. We split the cells from the same culture
into two equal portions. One portion was treated with the
impermeable Rh-6-(F-araNAD) and then the cells were lysed
Raji cells showed strong plasma membrane fluorescent
labeling by SR101−F-araNMN similar to RA-treated HL-60
cells (Figure 6A). To better visualize intracellular labeling, we
first blocked plasma membrane CD38 with the nonfluorescent,
cell impermeable probe, 6-alkyne-F-araNAD. Then, the cells
were incubated with SR101−F-araNMN. We also used DAPI
to stain the nucleus. Very weak fluorescence was observed in
the nucleus and most of the label localized in the cytoplasm
E
dx.doi.org/10.1021/ja411046j | J. Am. Chem. Soc. XXXX, XXX, XXX−XXX

Journal of the American Chemical Society
Article
Figure 7. (A) Confocal image of K562 cells with SR101−F-araNMN
labeling. Confocal microscope settings: laser power: 4.5%, pinhole: 1.1
airy units, master gain for PMT: 803. (B) K562 in-gel fluorescence
analysis: lanes 1, live cell labeling with SR101−F-araNMN followed by
in-gel fluorescence; lanes 2, whole cell lysate was obtained first
followed by labeling with SR101−F-araNMN (labeling all catalytically
active CD38); lanes 3, whole cell lysate without CD38 probe. Ladder
is on the left, listed first. The full gel images are shown in Figure S7 in
SI. (C) Western blot analysis for detection of CD38 in all cell lines.
Figure 6. Labeling of CD38 in Raji cells. (A) Confocal image of Raji
cells labeled with SR101−F-araNMN. Confocal microscope settings
for (A): laser power: 5.5%, pinhole: 1.1 airy units, master gain for
PMT: 866. (B) Confocal image of Raji cells blocked with 6-alkyne-(F-
araNAD) then labeled with SR101−F-araNMN (visualization of
intracellular CD38 only). DAPI (blue) staining dsDNA showing
nucleus. Confocal microscope settings for (B): laser power: 10%,
pinhole: 1.1 airy units, master gain for PMT: 866. An increase in laser
power and master gain was necessary in order to have enough
fluorescence emission to see the signal. This indicates a low amount of
intracellular active CD38. (C) In-gel fluorescence analysis: lanes 1, live
cell labeling with Rh-6-(F-araNAD) followed by in-gel fluorescence
(labeling plasma membrane CD38 only); lanes 2, whole cell lysate was
obtained first followed by labeling with Rh-6-(F-araNAD) (labeling all
catalytically active CD38); lanes 3, whole cell lysate without CD38
probes. Ladder is on the left, listed first. The full gel images are shown
in Figure S7 in SI.
this possibility, we further performed Western blot analysis on
whole cell lysate to detect CD38. The Western blot again
showed that K562 cells did not express CD38, while CD38 was
detected in RA-treated HL-60 and Raji whole cell lysates, along
with low expression in untreated HL-60 cells (Figure 7c).
Therefore, our data collectively demonstrated that K562 cells
did not express CD38.
DISCUSSION
■
We developed a cell-permeable CD38 probe, SR101−F-
araNMN, by choosing a more cell-permeable fluorescent dye
(SR101) and by using F-araNMN instead of F-araNAD. Using
purified CD38, SR101−F-araNMN could label wild type CD38
but not the catalytic mutant of CD38, confirming that it was an
activity-based probe. Using three human blood cancer cell lines,
we further confirmed that SR101−F-araNMN was cell
permeable and was able to specifically label CD38. Combined
with the previously developed impermeable CD38 probes, Rh-
6-(F-araNAD) and 6-alkyne-F-araNAD, we were able to better
visualize intracellular CD38 in these cell lines and obtained
CD38 localization information that could correct several
misconceptions about the intracellular distribution, which are
discussed below.
when methanol was used to wash off the probe (Figure 6B).
Similar results were obtained when we used PFA and methanol
to treat cells (Figure S4 in SI). We again used the in-gel
fluorescence analysis, which showed essentially the same
labeling intensities in the live cell labeling and the whole cell
lysate labeling with Rh-6-(F-araNAD), after correcting for
protein loading using the Coomassie blue staining (Figure 6C).
This result further supported the previous confocal imaging
results with SR101−F-araNMN and suggested that the amount
of intracellular CD38 was very little. Also, quantification using
6-alkyne-F-araNAD and SR101−F-araNMN as described above
for HL-60 cells, revealed that about 4.2% of catalytically active
CD38 in Raji cells is intracellular (Figure S3 in SI).
As for the K562 cells, no SR101−F-araNMN fluorescent
labeling was detected from confocal imaging (Figure 7A).
Consistent with this, labeling of cell lysate with SR101−F-
araNMN also failed to detect the presence of CD38 (Figure
7B). This result was contrary to the reported result showing
that K562 had only intracellular CD38. One possibility for the
discrepancy was that intracellular CD38 was present but that it
was not catalytically active for unknown reasons. To rule out
CD38 Is Highly Concentrated in the Plasma Mem-
branes of Raji and RA-Treated HL-60 Cells. It was reported
that CD38 was present mainly on the plasma membrane with
the catalytic domain facing outside.22 Consistent with this,
when we labeled CD38 in RA-treated HL-60 cells or untreated
Raji cells with SR101−F-araNMN, the strongest fluorescence
resided on the plasma membrane based on confocal imaging.
When limiting the fluorescence detection to intracellular CD38,
HL-60 cells showed nearly a 2.5-fold increase of intracellular
CD38 level upon RA treatment. However, the CD38
F
dx.doi.org/10.1021/ja411046j | J. Am. Chem. Soc. XXXX, XXX, XXX−XXX

Journal of the American Chemical Society
Article
expression on the plasma membrane had a much greater
increase based upon fluorescence detected. Additionally, our
probe was activity-based; therefore, the intracellular CD38 we
detected in RA-treated HL-60 and untreated Raji cells should
be catalytically active. As a consequence, if the intracellular
CD38 had access to cellular NAD, it might lead to decreased
intracellular NAD levels. On the basis of confocal imaging,
intracellular CD38 was found to be present in punctate bodies
in the cytosol, which were likely membrane organelles, such as
ER, Golgi, or mitochondria. At this point, we do not know the
exact identities of the intracellular organelles that contain
CD38, nor do we know the topology of CD38 (facing the
cytosol or the matrix) on these intracellular organelles.
Raji and RA-Treated HL-60 Cell Nuclei Have Little
CD38. A previous report on CD38 localization used confocal
immunofluorescence to show the presence of CD38 primarily
within the nucleus of Raji cells.15 In fact, it was shown that
CD38 colocalized in nuclear Cajal bodies, which are small
subnuclear membraneless organelles present either free in the
nucleoplasm and/or physically associated to specific regions of
chromatin. However, using the activity-based probes for CD38,
we found very little CD38 labeled within the nucleus, and
certainly no concentrated areas of fluorescence that would
represent active CD38 in Cajal bodies. In fact, the nucleus had
the weakest CD38 labeling in both Raji cells and RA-treated
HL-60 cells. Although we could not rule out the presence of
catalytic inactive CD38, the results obtained with the K562 cells
suggested that this was unlikely (discussed below). Our results
thus suggested that very little CD38 is present in the nuclei of
Raji and RA-treated HL-60 cells.
K562 Cells Do Not Have Detectable CD38 Expression.
It was reported on the basis of immunofluorescence that K562
cells express intracellular CD38 but no plasma membrane
CD38.15 When labeling K562 cells with SR101−F-araNMN, we
saw no fluorescence from confocal microscopy imaging on
either the plasma membrane or within the cell. To rule out that
SR101−F-araNMN was not permeable to K562 cells, we also
labeled K562 whole cell lysates with SR101−F-araNMN. No
fluorescent CD38 band was observed on SDS-PAGE gel,
consistent with the confocal imaging results. This result was
contrary to the reported intracellular only localization of CD38.
To resolve the conflicting observations, we first considered the
possibility that K562 cells had CD38 but that CD38 was
catalytically inactive; and thus, CD38 could not be labeled with
our probe. However, Western blot showed that K562 cells had
no detectable CD38 expression. Therefore, the labeling result
with SR101−F-araNMN in K562 cells was reliable and revealed
the absence of CD38 in K562 cells.
Antibodies vs Activity-Based Probes for Detecting
CD38 Intracellular Localization. The most commonly used
methods to detect CD38 cellular localization are confocal
microscopy coupled with immunofluorescence or GFP fusion.
Using GFP fusion proteins requires the overexpression of the
fusion proteins, which may cause artifacts due to the
overexpression or the fusion. Immunofluorescence has the
advantages of detecting endogenous proteins but it requires the
use of antibodies. CD38 antibodies are convenient tools to use
in most applications. However, for detecting the intracellular
localization of CD38, antibodies have some limitations. First,
antibodies are not cell permeable; thus, the cells have to be
fixed and permeablized before labeling with antibodies. Second,
washing conditions have to be controlled well to wash away
unbound antibodies but not antibodies that bind to the protein
target. The potential problems can be fixed but require extra
efforts to optimize experimental conditions. The reported
prominent nuclear CD38 in Raji and K562 cells is possibly a
false positive caused by insufficient washing, which could have
been avoided if proper negative and positive controls were used
to help find the optimal washing conditions.15 When inspecting
the data on the immunofluorescence detection of CD38 in Raji
and K562 cells, we noticed that there were no negative or
positive controls provided. Nuclear membrane localization was
also reported in other cell types.20,31 Immunofluorescence was
the major method used in these studies reporting the nuclear
membrane localization of CD38. These studies may also suffer
from similar problems. Compared with immunofluorescence,
our permeable CD38 probe has certain advantages that can
complement the use of antibodies. First, the probe identifies
catalytically active CD38 because labeling proceeds in an
activity-dependent way while immunofluorescence can be used
to reveal total protein levels. Second, the probe is cell
permeable and thus does not require detergent permeabiliza-
tion of cells. Thus, the labeling can be done under less
perturbing conditions. Unfortunately, we had to fix cells to
wash away excess probe molecule. Optimization of the probes
to allow aqueous buffer wash will be required for true live-cell
imaging. However, because it forms a rather stable covalent
linkage with CD38, the washing step can be done very
extensively to make sure all excess CD38 probes are washed
away. This way, fewer false positives will be observed. This is
probably an important feature that allowed us to conclude that
there is very little intracellular CD38. The availability of both a
permeable and impermeable CD38 probe greatly facilitated the
quantification of intracellular versus cell surface CD38, which is
otherwise not easy to accurately quantify.
CONCLUSION
■
NAD is increasingly recognized as an important signaling
molecule that regulates many physiologically important
processes. Understanding the function of NAD-metabolizing
enzymes, such as CD38, is thus important and can impact a
variety of different areas. Ensuring proper cellular function
requires the spatial distribution of different proteins to be
delicately regulated. In the case of CD38, localization is
similarly important to its biological function. Investigating its
correct cellular localization will help to understand several
unaddressed questions about the function of CD38. Early
studies on CD38 focused on its ability to make cADPR and
NAADP, which are considered important second messengers
capable of releasing intracellular stored calcium. However, in
vitro, the most efficient activity of CD38 is the hydrolysis of
NAD, with kcat of 96 s−1 and Km of 16 μM.32 If CD38 is present
in the nucleus as previously reported, it will likely deplete
nuclear NAD. Using the permeable and impermeable CD38
probes, we found that the nucleus actually contained the least
amount of fluorescence, which we interpreted as the lack of
CD38 in the nucleus. Considering the efficient NAD hydrolysis
activity, we think the lack of nuclear CD38 actually makes
sense; otherwise, intracellular NAD may be depleted. CD38-
catalyzed formation of cADPR accounted for less than 2% of
the total product. If this activity is physiologically important, it
is possible that a certain intracellular pool of CD38 may have
higher cyclase activity. CD38-catalyzed formation of NAADP in
vitro requires low pH and high concentration of nicotinic acid.
The low pH condition is only possible in certain acidic
organelles, such as the lysosome. Thus, investigating the exact
G
dx.doi.org/10.1021/ja411046j | J. Am. Chem. Soc. XXXX, XXX, XXX−XXX

Journal of the American Chemical Society
Article
(16) Orciani, M.; Trubiani, O.; Guarnieri, S.; Ferrero, E.; Di Primio,
R. J. Cell Biochem. 2008, 105, 905.
(17) Ceni, C.; Pochon, N.; Brun, V.; Muller-Steffner, H.; Andrieux,
A.; Grunwald, D.; Schuber, F.; De Waard, M.; Lund, F.; Villaz, M.;
Moutin, M. J. Biochem. J. 2003, 370, 175.
(18) Reinherz, E. L.; Kung, P. C.; Goldstein, G.; Levey, R. H.;
Schlossman, S. F. Proc. Natl. Acad. Sci. U.S.A. 1980, 77, 1588.
(19) Yamada, M.; Mizuguchi, M.; Otsuka, N.; Ikeda, K.; Takahashi,
H. Brain Res. 1997, 756, 52.
(20) Adebanjo, O. A.; Anandatheerthavarada, H. K.; Koval, A. P.;
Moonga, B. S.; Biswas, G.; Sun, L.; Sodam, B. R.; Bevis, P. J.; Huang,
C. L.; Epstein, S.; Lai, F. A.; Avadhani, N. G.; Zaidi, M. Nat. Cell Biol.
1999, 1, 409.
(21) Zhao, Y. J.; Lam, C. M.; Lee, H. C. Sci. Signal. 2012, 5, ra67.
(22) Moreno-Garcia, M. E.; Sumoza-Toledo, A.; Lund, F. E.; Santos-
Argumedo, L. Mol. Immunol. 2005, 42, 703.
(23) Jiang, H.; Congleton, J.; Liu, Q.; Merchant, P.; Malavasi, F.; Lee,
H. C.; Hao, Q.; Yen, A.; Lin, H. J. Am. Chem. Soc. 2009, 131, 1658.
(24) Liu, Q.; Kriksunov, I. A.; Graeff, R.; Munshi, C.; Lee, H. C.;
Hao, Q. Structure 2005, 13, 1331.
intracellular localization of CD38 will likely provide insights
into the physiological relevance of different enzymatic activities
of CD38. The activity-based probes we developed will allow the
determination of intracellular localization of CD38 in different
cells and thus enable a better understanding of the physiological
function of CD38.
ASSOCIATED CONTENT
* Supporting Information
■
S
Details of experimental procedures, acquired and measured
NMR, and LC−MS spectra for the synthesis of the CD38
probe, SR101−F-araNMN. This material is available free of
charge via the Internet at http://pubs.acs.org.
AUTHOR INFORMATION
Corresponding Author
hl379@cornell.edu
■
Notes
(25) Munshi, C.; Aarhus, R.; Graeff, R.; Walseth, T. F.; Levitt, D.;
Lee, H. C. J. Biol. Chem. 2000, 275, 21566.
The authors declare no competing financial interest.
(26) Kolb, H. C.; Finn, M. G.; Sharpless, K. B. Angew. Chem., Int. Ed.
2001, 40, 2004.
(27) Lamkin, T. J.; Chin, V.; Varvayanis, S.; Smith, J. L.; Sramkoski,
R. M.; Jacobberger, J. W.; Yen, A. J. Cell Biochem. 2006, 97, 1328.
(28) Schnell, U.; Dijk, F.; Sjollema, K. A.; Giepmans, B. N. Nat.
Methods 2012, 9, 152.
ACKNOWLEDGMENTS
■
This work is supported by NIH R01GM086703 (H.L.) and
R01CA1525870 (A.Y.). The confocal imaging was enabled by
the Cornell University Biotechnology Resource Center that is
supported by NIH 1S10RR025502-01 for all data collected on
the Zeiss LSM 710 Confocal.
(29) Rockstroh, M.; Muller, S.; Jende, C.; Kerzhner, A.; Bergen, M.;
Tomm, J. J. Integr. Omics 2011, 1, 135.
(30) Cox, B.; Emili, A. Nat. Protoc. 2006, 1, 1872.
REFERENCES
(1) Houtkooper, R. H.; Canto,
Rev. 2010, 31, 194.
(2) Lin, H. Org. Biomol. Chem. 2007, 5, 2541.
■
(31) Khoo, K. M.; Han, M. K.; Park, J. B.; Chae, S. W.; Kim, U. H.;
Lee, H. C.; Bay, B. H.; Chang, C. F. J. Biol. Chem. 2000, 275, 24807.
(32) Sauve, A. A.; Munshi, C.; Lee, H. C.; Schramm, V. L.
Biochemistry 1998, 37, 13239.
́
C.; Wanders, R. J.; Auwerx, J. Endocr.
(3) Partida-Sanchez, S.; Cockayne, D. A.; Monard, S.; Jacobson, E. L.;
Oppenheimer, N.; Garvy, B.; Kusser, K.; Goodrich, S.; Howard, M.;
Harmsen, A.; Randall, T. D.; Lund, F. E. Nat. Med. 2001, 7, 1209.
(4) Jin, D.; Liu, H. X.; Hirai, H.; Torashima, T.; Nagai, T.; Lopatina,
O.; Shnayder, N. A.; Yamada, K.; Noda, M.; Seike, T.; Fujita, K.;
Takasawa, S.; Yokoyama, S.; Koizumi, K.; Shiraishi, Y.; Tanaka, S.;
Hashii, M.; Yoshihara, T.; Higashida, K.; Islam, M. S.; Yamada, N.;
Hayashi, K.; Noguchi, N.; Kato, I.; Okamoto, H.; Matsushima, A.;
Salmina, A.; Munesue, T.; Shimizu, N.; Mochida, S.; Asano, M.;
Higashida, H. Nature 2007, 446, 41.
(5) D’Arena, G.; Musto, P.; Cascavilla, N.; Dell’Olio, M.; Di Renzo,
N.; Perla, G.; Savino, L.; Carotenuto, M. Leuk. Lymphoma 2001, 42,
109.
(6) States, D. J.; Walseth, T. F.; Lee, H. C. Trends Biochem. Sci. 1992,
17, 495.
(7) Howard, M.; Grimaldi, J. C.; Bazan, J. F.; Lund, F. E.; Santos-
Argumedo, L.; Parkhouse, R. M.; Walseth, T. F.; Lee, H. C. Science
1993, 262, 1056.
(8) Kim, H.; Jacobson, E. L.; Jacobson, M. K. Science 1993, 261,
1330.
(9) Lee, H. C.; Zocchi, E.; Guida, L.; Franco, L.; Benatti, U.; De
Flora, A. Biochem. Biophys. Res. Commun. 1993, 191, 639.
(10) Takasawa, S.; Tohgo, A.; Noguchi, N.; Koguma, T.; Nata, K.;
Sugimoto, T.; Yonekura, H.; Okamoto, H. J. Biol. Chem. 1993, 268,
26052.
(11) Chini, E. N.; Chini, C. C.; Kato, I.; Takasawa, S.; Okamoto, H.
Biochem. J. 2002, 362, 125.
(12) Zubiaur, M.; Fernandez, O.; Ferrero, E.; Salmeron, J.; Malissen,
B.; Malavasi, F.; Sancho, J. J. Biol. Chem. 2002, 277, 13.
(13) Khoo, K. M.; Chang, C. F. Int. J. Biochem. Cell Biol. 2002, 34, 43.
(14) Yalcintepe, L.; Albeniz, I.; Adin-Cinar, S.; Tiryaki, D.; Bermek,
E.; Graeff, R. M.; Lee, H. C. Exp. Cell Res. 2005, 303, 14.
(15) Trubiani, O.; Guarnieri, S.; Eleuterio, E.; Di Giuseppe, F.;
Orciani, M.; Angelucci, S.; Di Primio, R. J. Cell Biochem. 2008, 103,
1294.
H
dx.doi.org/10.1021/ja411046j | J. Am. Chem. Soc. XXXX, XXX, XXX−XXX